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Research Branch 
Technical Bulletin 1997-01 



Monograph 

on 
Downey Mildew 

of 
Crucifers 



Saskatoon Research Centre 



Canada 



Cover illustration 

The images represent the Research Branch's objective: 
to improve the long-term competitiveness of the Canadian 
agri-food sector through the development and transfer of new 
technologies. 

Illustration de la couverture 

Les dessins illustrent l'objectif de la Direction generate de la 
recherche : ameliorer la competitivite a long terme du secteur 
agro-alimentaire canadien grace a la mise au point et au transfert 
de nouvelles technologies. 



® 



Monograph 

on 
Downy Mildew 

of 
Crucifers 



G.S. Saharan 1 , P.R. Verma 2 and N.I. Nashaat 3 



1. Department of Plant Pathology, CCS. Haryana Agricultural University, Hisar, 125004, 
Haryana, India 

2. P.R. Verma, Agriculture and Agri-Food Canada, Saskatoon Research Centre, 107 Science 
Place, Saskatoon, Saskatchewan, S7N 0X2, Canada 

3. N.I. Nashaat, Crop and Disease Management Department, Institute of Arable Crops- 
Research Rothamsted, Harpenden, Hertfordshire, AL5 2JQ, United Kingdom 



Saskatoon Research Centre 

Technical Bulletin 1997-01 

Research Branch 

Agriculture and Agri-Food Canada 

1997 



Copies of this publication are available from 

Director 

Saskatoon Research Centre 

Research Branch, Agriculture and Agri-Food Canada 

107 Science Place 

Saskatoon, Saskatchewan, Canada S7N 0X2 



@ Minister of Supply and Services Canada 1997 
Cat. No. A5A-13/1997-01E 
ISBN 0-662-25744-8 



CONTENTS 

FOREWORD 1 

PREFACE 2 

ACKNOWLEDGEMENTS 3 

1. INTRODUCTION 4 

2. THE DISEASE 4 

a. Geographical distribution 5 

b. Economic importance 5 

i) Brassica oilseeds 5 

ii) Brassica vegetables 5 

c. Host range 9 

d. Symptoms 9 

i) Brassica oilseeds 9 

ii) Brassica vegetables 15 

iii) Broccoli 16 

iv) Wallflower 17 

v) Stock ." 17 

e. Disease assessment 17 

3. THE PATHOGEN 18 

a. Taxonomy and morphology 20 

b. Reproduction and reproductive structures 20 

i) Asexual phase 20 

ii) Sexual phase 24 

c. Electron microscopy and ultra structures 25 

i) Host penetration 25 

ii) Haustorium development 29 

iii) The host - pathogen interface 31 

iv) Conidiophore development 38 

v) Conidiophore growth 41 

vi) Conidial formation 45 

vii) Host response 45 

viii) Cytology and genetics 45 

d. Physiologic specialization (Pathogenic variability) 51 

e. Heterothallism and homothallism 60 

f. Perpetuation 63 

i) Mycelium 63 



ii) Conidia 63 

iii) Oospores 64 

iv) Axenic culture . 64 

g. Conidial discharge 65 

h. Conidial germination 69 

i. Oospore germination 69 

4. INFECTION AND PATHOGENESIS 69 

5. DISEASE CYCLE 74 

6. EPIDEMIOLOGY 76 

a. Disease development in relation to temperature, humidity, rainfall, 

and leaf wetness 76 

b. Disease development in relation to planting time 83 

c. Disease development in relation to host nutrition 83 

d. Disease interaction with insecticidal sprays 88 

7. MIXED INFECTIONS AND ASSOCIATION WITH WHITE RUST .... 88 

8. BIOCHEMISTRY OF THE HOST PATHOGEN INTERACTION 92 

a. Metabolic changes 92 

b. Role of natural biochemical compounds 98 

9. RESISTANCE 99 

a. Mechanism of host resistance 99 

b. Host-pathogen recognition system 106 

c. Systemic acquired resistance 106 

d. Genetics of host-pathogen relationship 106 

e. Biochemical basis of resistance 109 

f. Lignification of host cells Ill 

g. Sources of resistance 114 

10. BREEDING FOR DISEASE RESISTANCE 114 

11. DEVELOPMENT OF RESISTANCE TO FUNGICIDES 115 

12. LABORATORY AND FIELD TECHNIQUES AND BIOASSAYS 116 

a. Culturing of Peronospora parasitica 116 

b. Maintenance of P. parasitica isolates 120 



c. Germplasm screening and evaluation 121 

d. Preservation of P. parasitica 124 

e. Artificial inoculation of excised cotyledons 124 

f. Propagation of P. parasitica on cotyledons or true leaves of Japanese 
radish seedlings 125 

g. Laboratory tests of fungicides 125 

h. Fungicide resistance assay 126 

i. Measuring systemic infection by the downy mildew pathogen 127 

j. Methods of breeding for multiple disease resistance 127 

k. Heterothallism and homothallism 130 

1. Seed-borne nature of P. parasitica 132 

m. Conidial germination 133 

n. Sporulation 133 

o. Discharge of Conidia 133 

13. DISEASE MANAGEMENT 134 

a. Cultural practices 134 

b. Seed treatment 134 

c. Soil treatment 135 

d. Compost treatment 135 

e. Foliar spray of fungicides 136 

i) Brassica vegetables 136 

ii) Brassica oilseeds 137 

f. Biological control 138 

i) Plant extracts as fungitoxicant 138 

ii) Antagonists for biocontrol 142 

g. Host resistance 142 

h. Integrated disease management 142 

14. FUTURE STRATEGIES AND PRIORITIES OF DOWNY MILDEW 
DISEASE MANAGEMENT 145 

a. Disease epidemiology 145 

b. Physiological specialization 145 

c. Genetics of resistance 145 

d. Molecular aspects 147 

e. Biochemical aspects of resistance 150 

f. Disease management 151 

15. REFERENCES 151 

16. SUBJECT INDEX 180 



Digitized by the Internet Archive 

in 2013 



http://archive.org/details/monographondown199701cana 



FOREWORD 

Crucifer oilseed, vegetables and fodder crops represent an increasing percentage of the 
agricultural economies of many nations occupying important niches in temperate, cool 
temperate, continental and subtropical tropical regions of the world. 

Vegetable crucifers continue to be a major source of vitamins, fiber, minerals and proteins 
in the human diet, while crucifer seed oil consumption and industrial utilization increases 
annually. Substantial efforts are underway to improve the quality of crucifer seed oils through 
genetic engineering and traditional breeding. With this global expansion in crucifer crop 
production comes the increasing potential for severe losses due to damage and disease of insect 
pests and pathogens. This monograph on the downy mildews of crucifers, therefore, is a most 
timely contribution to our knowledge of a most important global pathogen of crucifers, 
Peronospora parasitica (Pers. ex Fr.) Fr. 

One might believe that at this time at the dawn of the 'information age' in which ready 
access to current information and newly emerging knowledge is increasingly available to all 
through the internet that there would be little need to bring together all of the relevant 
information into the format of a printed monograph. I would assert that quite the contrary is the 
case. This Monograph on Downy Mildew of Crucifers by its inclusiveness, carefully crafted 
organization and thorough documentation of the existing research and reported literature 
contextualized to be relevant to both the researcher and the practitioner is a most valuable 
'benchmark' publication. In this age of information, this monograph provides the much needed 
background references and information together with the most current insights and 
methodologies as to ensure its place as a central document for students, teachers, researchers 
and practitioners investigating this organism and its hosts. 

The authors, G. S. Saharan, P. R. Verma and N. I. Nashaat, bring their lifelong 
professional interest and expertise to the presentation of this treatise. Together, they have 
crafted a most useful document in which a wide range of information is logically organized and 
easily accessed. This is an important contribution in a series including Alternaria diseases and 
white rust of crucifers that have appeared as Technical Bulletins from the Research Branch of 
Agriculture and Agri-Food Canada. 




\ o— & tV-v^O JUu^ j 



-\ 



April, 1997 Paul H. Williams 

Atwood Distinguished Professor of Plant 

Pathology 
University of Wisconsin-Madison 
Madison, WI USA 



PREFACE 

This treatise on downy mildew of crucifers was compiled as the natural extension 
of a comprehensive literature search published in the form of a bibliography in 1994. 
Downy mildew is amongst the major devastating diseases of crucifers worldwide. The 
most common hosts of this disease are high quality edible oil crops (rapeseed-mustard, 
canola and other rapes), industrial oil crop (crambe and other rapes), common 
vegetables (cabbage, cauliflower, radish, kohlrabi, broccoli, brussels sprouts, kales, and 
other Brassica vegetables), and ornamental plants (wallflower and stocks). Weeds are 
also common hosts. 

For convenience of the readers, the information has been arranged into sections 
and subsections. The sections discuss subjects relating to: i) the disease, its symptoms 
on different hosts, geographic distribution, yield losses and disease assessment 
methods; ii) the pathogen's taxonomy, variability, sporulation, perpetuation and spore 
germination; iii) host-pathogen interactions in the form of seed infection, host range, 
disease cycle, process of infection and pathogenesis, epidemiology, fine structure, 
biochemical changes and biochemical compounds; iv) host defense mechanisms; v) 
techniques to study host-pathogen relationships; and vi) management practices related 
to cultural control, chemical control, biological control, host resistance and integrated 
disease management. To stimulate new ideas in downy mildew research, a section on 
future priorities has been included. 

We hope that this monograph on downy mildew will be useful to research 
scientists, teachers, extension specialists, students, industries and all others who are 
working with cruciferous crops and striving for crop improvement through disease 
management. 

G.S. Saharan 
Hisar, India 

P.R. Verma 
Saskatoon, Canada 

N.I. Nashaat 
Harpenden, U.K. 



ACKNOWLEDGEMENTS 

An undertaking of this nature cannot succeed without the help of many people. It 
is a pleasure to acknowledge those who have contributed. The original tracking down 
of many papers was done by Saskatoon Research Centre Librarians, Van Keane and 
Gail Charabin. Three other people deserve special acknowledgement: Ralph 
Underwood helped reproduce photographs from original journals and Ila Woroniuk and 
Jan Korven-Stott spent many hours typing, proof reading and preparing a final 
version. Any errors, either of commission or omission are, however, our responsibility. 
We hope they will be brought to our attention. 

During the preparation of this monograph, several of the second authors' 
colleagues, including Dr. PA. O'Sullivan as Director, Dr. R.K. Downey as Emeritus 
Scientist, and Dr. G.F.W. Rakow as Chairman of the Oilseed Section, at Saskatoon, 
have made valuable suggestions and have been very supportive of the work. The 
authors also thank Drs. KL. Bailey and L.J. Duczek, Research Scientists, Agriculture 
and Agri-Food Canada, Research Centre, Saskatoon for critically reviewing the 
manuscript. Here, we also wish to thank many people at Headquarters Library, 
Agriculture and Agri-Food Canada, Ottawa, for providing countless photocopies, and 
to the Translation Bureau, Department of Secretary of State for translation of a series 
of foreign language papers. 

The senior author also thanks Dr. J.B. Chowdhury, Vice-Chancellor, Dr. D.P. 
Singh, Dean, College of Agriculture and Dr. M.P. Srivastava, Head, Department of 
Plant Pathology, CCS. Haryana Agricultural University, Hisar, India, for permitting 
him to spend two months in the second author's laboratory to co-author this 
publication. 



1. INTRODUCTION 

The term "mildew" was first used in the United States to denote a wide group of 
Pparasitic fungi with little in common except their appearance as a white or lightly 
coloured delicate outgrowths caused by the proliferation and fructification of mycelium 
on the surface of green and necrotic plant tissues. Downy mildew quickly adapted to 
European conditions when Vine mildew was introduced from North America. Downy 
mildew or members of the family Peronosporacae are a distinctive group of obligate 
plant pathogens classified within the Mastigomycotina in the Oomycete order 
Peronosporales. In the family cruciferae, about 50 genera and more than 100 different 
species are susceptible to infection by downy mildew pathogen. Originally Gaumann 
(1918), on the basis of conidial measurements and cross inoculation tests, recognized 
52 species of Peronospora on crucifer hosts. Later studies by Yerkes and Shaw (1959) 
concluded that there are no reliable morphological criteria for distinguishing 
Peronospora isolates from different host species, and all collections of downy mildew 
from the cruciferae are currently grouped in the single aggregate species P. parasitica 
(Pers. ex Fr.) Fr. Constantinescu (1989) later proposed a new genus, Paraperonospora, 
to accommodate several species of Peronospora pathogenic on hosts in the family 
compositae. 

There are two different patterns of host colonization: systemic and localized. 
Systemic infection is characterized by colonization of leaves, stems and sometimes roots, 
mostly through the infection of the seedlings by primary inoculum. The symptoms vary 
from chlorotic discolouration to stunting and distortion of the whole plant. Localized 
infections are characterized by the occurrence of lesions on leaves, surrounded by a 
conspicuous characteristic white "down" on the abaxial surface (Lucas and Sherriff, 
1988). 



2. THE DISEASE 

It is commonly known as "mildew", "mould", "false oidium or mildew" (French), 
"Falscher-Mehltau" (German) and more commonly by the name of downy mildew. The 
disease is caused by the fungus Peronospora parasitica. 

The upper surface of affected young and older leaves have ill-defined, irregular, pale 
yellow necrotic lesions, whereas the lower surface is covered by white grey mycelium. 
The cotyledons and older leaves may be killed prematurely when single lesions coalesce 
to form large blotches. Attacked pods may be covered with angular brown lesions, or 
under high humidity, a sparse white-greyish mycelium may develop. Severe attacks 
may lead to premature ripening. Geographic distribution, economic importance, 
symptoms on various hosts, host range and disease assessment are discussed below. 



a. Geographical distribution 

Downy mildew on cultivated Brassica species and other cruciferous host species is 
prevalent in widely separated localities in numerous countries throughout the world 
(Channon, 1981; Verma et al., 1994). World records of P. parasitica causing downy 
mildew disease on crucifers are given in Table 1. The names of various hosts in this 
table are as reported in the original papers. 

b. Economic importance 

The economic importance of Peronospora parasitica (downy mildew) has been 
adequately documented over the years. This pathogen, alone or in combination with 
Albugo Candida (white rust), is responsible for causing severe losses in yield of several 
temperate and tropical Brassicaceae crops, particularly rapeseed and mustard. Yield 
losses due to downy mildew infection alone is very difficult to estimate, since in most 
cases it is always associated with white rust. 

i) Brassica oilseeds: Hypertrophied host tissues termed as staghead are often 
observed in association with a mixed infection of A. Candida and P. parasitica 
particularly at the flowering stage. Yield losses in B. rapa var. toria (Toria) due to such 
combined infections is estimated to be about 34%, when the average length of individual 
hypertrophied racemes is 10 cm (Kolte, 1985). The combined infection with both 
pathogens on B. juncea may cause 37-47% and 17-32% reduction in siliques formation 
and seed production respectively (Bains and Jhooty, 1979). Others have reported 23- 
55% yield loss in the same host species due to the mixed infection with both pathogens 
(Saharan, 1984, 1992a). Kolte (1985) suggested the following formula for estimating the 
yield loss due to infection with white rust or downy mildew alone, or for combined 
infections: Q = A-(BxC) x 100 

A 
where: Q = percentage yield 

A = average actual or expected yield of a healthy plant 
B = average or expected yield from the affected raceme, which is equal 
to the actual average yield from the corresponding length of the 
healthy raceme 
C = number of affected racemes per plant. 

ii) Brassica vegetables: During 1911-1912, downy mildew infection in 
cabbage near Lahore, Pakistan caused more than 50% yield loss (Butler, 1918). 
Vasileva (1976) reported that under favourable conditions, P. parasitica may infect 
up to 50-60% of cabbage seeds and reduces yield by 16-20%. 



Table 1. World records of Peronospora parasitica on crucifers (Verma et 
al., 1994) 





Recording 






Location 


Year 


Host 


Reference 


Argentina 


1939 


Cabbage, Radish, Swede 


Lindquist, 1946 


Australia 


1924 


Cauliflower, Cabbage 


Samuel, 1925 


Austria 


1969 


Cabbage 


Glaeser, 1970 




1987 


Radish, Chinese Cabbage 


Bedlan, 1987 




i989 


Cabbage 


Bedlan, 1989 


Bavaria 


1936 


Horseradish 


Boning, 1936 


Bermuda 


1939 


Stock 


Waterston, 1940 


Borneo 


1962 


Chinese Cabbage 


Anonymous, 1962 


Brazil 


1943 


Cabbage & Broccoli 


Viegas & Teixeira, 1943 


Britain (U.K.) 


1948 


Capsella-bursa-pastoris 


Foister, 1948 




1959 


Broccoli, Brussels sprouts, cabbage, 
Cauliflower, Kale, Kohl-rabi, Marrow- 
stem Kale, Rape, Turnips, Radish, 
Horseradish, Swede, Stock, 
Wallflower, Watercress 


Moore, 1959 


Brunei 


1981 


Crucifers 


Channon, 1981 


Bulgaria 


1979 


Turnip 


Khristov, 1979 


Canada 


1944 


Cauliflower 


Jones, 1944 




1961 


Rape, Crucifers 


Downey & Bolton, 1961 


Chile 


1960 


Crucifers 


Mujica & Vergara, 1960 


China 


1925 


Rape 


Porter, 1926 




1957 


Cabbage 


Pai, 1957 


Costa Rica 


1967 


Crucifers 


McGuire & Crandall, 1967 


Cuba 


1973 


Crucifers 


Fernandez, 1973 


Cypress 


1981 


Crucifers 


Channon, 1981 


Czechoslovakia 


1968 


Crucifers 


Rydl, 1968 


Denmark 


1924 


Crucifers 


Gram & Rostrup, 1924 




1949 


Stock 


Anonymous, 1949 


Dominica 


1972 


Crucifers 


Anonymous, 1972a 


Ethiopia 


1981 


Crucifers 


Channon, 1981 


Fiji 


1969 


Chinese Cabbage 


Anonymous, 1969 


Finland 


1981 


Crucifers 


Channon, 1981 


France 


1943 


B. napus, turnip, Camelina sativa, 
Sinapsis alba 


Darpoux, 1945 


Germany 


1938 


Colza 


Klemm, 1938 




1939 


Rape 


Raabe, 1939 




1955 


Cabbage 


Neumann, 1955 


Greece 


1981 


Crucifers 


Channon, 1981 


Guatemala 


1950 


Brassica spp. 


Muller, 1950 


Haiti 


1972 


Crucifers 


Anonymous, 1972a 


Holland 


1924 


Cabbage 


Thung, 1926a 


Hong Kong 


1962 


B. alboglabra 
Chinese Kale 


Johnston, 1963 



Hungary 


1957 


Stock 


Lehoczky, 1957 


Iberica (Spain) 


1924 


B. napus, B. oleracea 


Gonzalez, 1924 


India 


1918 


Brassica spp., Crucifers 


Butler, 1918 




1940 


B. campestris, B. napus, Radish 
Eruca sativa, Maledmia africana, 
Sisymbrium iris 


Thind, 1942 




1968 


Capsalla bursa - pastoris 


Rao, 1968 




1976 


Cardamine impatiens 


Sharma & Munjal, 1977 




1980 


Kohlrabi, Kale 


Puttoo & Choudhary, 1984 




1981 


B. pekinensis 


Karwasra & Saharan, 1982 




1982 


Cabbage 


Gupta & Choudhary, 1987 


Iran 


1989 


Radish 


Etebarian, 1989 


Iraq 


1981 


Crucifers 


Channon, 1981 


Ireland 


1970 


Cauliflower 


McKee, 1971 


Israel 


1953 


Cabbage, Cauliflower 


Peleg, 1953 


Italy 


1961 


Crucifers 


Ciferri, 1961 


Jamaica 


1967 


B. oleracea 


Leather, 1967 


Japan 


1934 


Brassica spp., Crucifers 


Hiura & Kanegae, 1934 


Kampuchea 


1969 


Crucifers 


Soonthronpoct, 1969 


Kenya 


1957 


Kale 


Anonymous, 1957 


Korea 


1972 


Crucifers 


Anonymous, 1972b 




1981 


Chinese Cabbage 


So et al., 1981 


Libya 


1981 


Crucifers 


Channon, 1981 


Ludlow 


1929 


Swedes 


Preston, 1929 


Malawi 


1972 


Crucifers 


Peregrine & Siddiqi, 1972 


Malaysia 


1949 


B. rapa 


Mcintosh, 1951 


Malta 


1981 


Crucifers 


Channon, 1981 


Mauritius 


1950 


Cabbage 


Orian, 1951 


Mexico 


1983 


Rapeseed 


Ponce & Mendoza, 1983 


Montpellier 


1941 


Stock 


Kuhnholtz & Gastaud, 1943 


Moravea 


1928 


Radish 


Baudys, 1928 


Morocco 


1981 


Crucifers 


Channon, 1981 


Mozambique 


1948 


Crucifers, Cabbage 


De Carvalho, 1948 


Nepal 


1966 


Crucifers 


Bhatt, 1966 


Netherlands 


1926 


Crucifers, Cabbage 


Thung, 1926b 


New South Wales 


1938 


Cauliflower, Mustard, Kohlrabi, Turnip 


Anonymous, 1938 


(Australia) 


1955 


Cabbage 


Anonymous, 1955 




1959 


Brussels Sprouts 


Anonymous, 1960b 




1966 


Stock 


Bertus, 1968 


New Zealand 


1963 


Crucifers 


Channon, 1981 


Norway 


1969 


Cabbage, Chinese Cabbage, Kohlrabi, 
Kale, Red Cabbage, Rape, Turnip, 
Radish 


Semb, 1969 


Palestine 


1935 


Cauliflower 


Rayss, 1938; Chorin, 1946 


Panama 


1967 


Crucifers 


McGuire & Crandall, 1967 


Pakistan 


1969 


Brassica, Crucifers 


Perwaiz et al., 1969 


Papua New Guinea 


1981 


Crucifers 


Channon, 1981 


Philippines 


1925 


B. juncea, B. pekinensis 


Ocfemia, 1925 


Poland 


1970 


Crucifers 


Zarzycka, 1970 


Portugal 


1953 


Cabbage 


Da Costa & Da Camara, 1953 



Puerto Rico 


1972 


Crucifers 


Channon, 1981 


Queensland 


1948 


Caronopus didymus 


Langdon, 1948 


Romania 


1930 


B. napus, B. nigra, Capsella 


Savulescu and Rayss, 1930 




1948 


Wallflower 


Savulescu, 1948 


Russia 


1989 


Radish 


Timina et al., 1989 


Sabah 


1962 


Crucifers 


Anonymous, 1962 


Samoa 


1975 


Crucifers 


Firman, 1975 


Saxony 


1927 


Wallflower, Stocks 


Wiese, 1927 


South Africa 


1934 


Cabbage, Cauliflower, Turnips, 
Radish, Kohlrabi 


Dippenaar, 1934 


Spain 


1924 


B. napus, B. oleracea 


Gonzalez, 1924 


Sri Lanka 


1932 


Crucifers 


Park, 1932 


Sweden 


1931 


Radish 


Hammarlund, 1931 




1944 


Colza, White mustard 


Bjorling, 1944 




1952 


Camelina sativa 


Borg, 1952 


Switzerland 


1923 


Brassica spp., Crucifers 


Gaumann, 1923 


Taiwan 


1961 


Crucifers 


Lo, 1961 


Tanzania 


1981 


Crucifers 


Channon, 1981 


Thailand 


1962 


Crucifers 


Chandrasrikul, 1962 


Trinidad and 








Tobago 


1922 


Cabbage 


Stell, 1922 


Turkey 


1981 


Crucifers 


Channon, 1981 


Uganda 


1981 


Crucifers 


Channon, 1981 


USA 


1883 


Brassica spp., Crucifers 


Farlow, 1883 




1889 


Sisymbrium spp., Lepidium 


Swingle, 1890 




1903 


Cauliflower 


Schrenk, 1905 




1918 


Turnip 


Gardner, 1920 




1923 


Cabbage 


Harter & Zones, 1923 




1927 


Watercress 


Davis, 1929 




1932 


Cabbage, Crucifers, Brassica spp. 


Weber, 1932 




1940 


Horseradish 


Kadow & Anderson, 1940 




1942 


Cabbage 


Snyder & Baker, 1943 




1954 


Radish 


Thompson & Decker, 1955 




1960 


Brassica spp., Crucifers 


Anonymous, 1960a 


Uruguay 


1955 


Crucifers 


Koch & Boasse, 1955 


USSR 


1955 


Cabbage 


Pimenova & Maslennikov, 1955 


Venezuela 


1981 


Crucifers 


Channon, 1981 


Vietnam 


1966 


Crucifers 


My, 1966 


Yugoslavia 


1954 


Cabbage 


Sutic & Khjajic, 1954 




1961 


Horseradish 


Macek, 1961 



Downy mildew disease can significantly affect the yield and developmental 
characters of radish (Achar, 1992b). Variables affected are the size and weight of 
silique, number of silique/plant, number of seeds/silique and weight of seeds. Seed yield 
loss can be as high as 58%. Infection also adversely affects the size and weight of roots. 
Disease loss assessment have been estimated according to the following equation: 



{ Mean yield of healthy plants - Mean yield of diseased plants} 

Yield loss (%) = X 100 

Mean yield of healthy plants 

c. Host range 

Few detailed studies have been made to determine the extent of the host ranges 
affected by downy mildew. In earlier work these fungi were inoculated on mature host 
tissues and then scored for the presence or absence of disease symptoms. The results 
suggested that downy mildew fungi had a very restricted host range (Gaumann, 1918). 
Peronospora on crucifers was originally examined with this assumption in mind. As a 
result a large number of species were created, mostly based on their occurrence on a 
particular crucifer genus. This process continued until Yerkes and Shaw (1959) called 
attention to the remarkable morphological similarity of the Peronospora species which 
attack crucifers. Following an extensive biometric study they reduced over eighty 
species names to synonymy with P. parasitica (Pers. ex Fr.) Fr. More recent work, 
involving examination of the reactions of crucifers seedlings to infection, has led to an 
even wider host range being established for P. parasitica (Foster, 1947a; Davison, 1967; 
McMeekin, 1969). Wide variation can be encountered in the reaction of seedlings of 
different crucifer species to the isolates of Peronospora from Brassica and Raphanus 
(Tables 2, 3), but the pathogen can grow well enough to sporulate on several species, 
apart from the original host (Dickinson and Greenhalgh, 1977). P. parasitica can infect 
a wide range of Brassica and other cruciferous species. Gaumann (1923) listed over 80 
cruciferous species as susceptible to infection by the numerous species of Peronospora 
which are now regarded as all being P. parasitica. Among the common hosts of 
economic importance are rapeseed-mustard, cabbage, Chinese cabbage, cauliflower, 
broccoli, brussels sprouts, marrow, stem kale, kohl rabi, turnip, turnip rape, swede, 
oilseed rape (canola), mustard, radish, horseradish, collards, rutabaga, watercress, stock 
and wallflower (Channon, 1981; Verma, et al., 1994; Nashaat, 1997). Apart from these, 
the inventory of hosts reported to be infected by P. parasitica are given in Table 4. 

d. Symptoms 

Downy mildew (P. parasitica) is the most frequently recorded disease on 
horticultural and agricultural members of the genus Brassica. The disease mainly 
affects young plants that may, in severe cases, be stunted or killed. Infection at later 
stages results in the debilitation and reduction in performance and quality of the host 
plant. 

i) Brassica oilseeds: Rapeseed-mustard : The disease appears on all above- 
ground plant parts but its symptoms are usually more conspicuous on leaves, stems and 
inflorescences. At the seedling stage on cotyledons and the first few true leaves, small 
angular translucent light-green lesions appear. These lesions later enlarge and develop 



10 

Table 2. Reaction of seedling cotyledons of members of the Cruciferae to 
inoculation with Brassica and Raphanus forms of P. parasitica. 
(Reactions were scored: 1 = no symptoms; 2 = necrotic flecking; 3 = 
extensive necrosis + slight sporulation; 4 = heavy sporulation; 1/2, 
etc., indicates intermediate reactions.) (Reprinted from C.H. 

Dickinson and J.R. Greenhalgh. 1977. Host range and taxonomy of 
Peronospora of crucifers. Trans. Brit. Mycol. Soc. 69: 111-116, by 
permission of the authors and the publisher British Mycological Society) 







Pathosren 


r 




Brassica 


Raphanus 


Host 




form 


form 


Brassica oleracea L. subsp. oleracea L. 


Wild cabbage 


3/4 


3 


B. oleracea L. 


Cultivated brassicas 


4 


3 


B. nigra (L.) Koch 


Black mustard 


1 


1 


B. juncea (L.) Czern 


Brown mustard 


3/4 


2 


B. rapa L. 


Turnip 


3 


3 


B. pekinensis (Lour.) Rupr. 


Chinese cabbage 


3 


2 


Sinapis alba L. 


White mustard 


2 


2 


Raphanus raphanistrum L. 


Wild radish 


2 


4 


R. maritimus Sm. 


Sea radish 


2 


4 


R. sativus L. 


Cultivated radish 


2 


4 


Crambe maritima L. 


Seakale 


1 


1 


Cakile maritima Scop. 


Sea rocket 


1 


1 


Lepidium sativum L. 


Garden cress 


1 


1 


Isatis tinctoria L. 


Woad 


2/4 


3 


Iberis amara L. 


Wild candytuft 


3/4 


3 


I. umbellata L. 


Garden candytuft 


3/4 


3 


/. sempervirens L. 


Perennial candytuft 


3 


3 


Thlaspi arvense L. 


Field pennycress 


2 


3 


T. rotundifolium (L.) Gaudin 


— 


2 


3 


Aethionema grandifloria R. Br. 


... 


1 


1 


Capsella bursa-pastoris (L.) Medic. 


Shepherd's purse 


1 


1 


Lunaria annua L. 


Honesty 


2 


1 


Alyssum saxatile L. 


Golden aylssum 


2 


2 


A. maritimum (L.) Lam. 


Sweet Alison 


2 


2 


Draba pyrenaica L. 


— 


1 


1 


Arabis alpina L. 


Alpine rock cress 


2 


1 


A. caerulea (All.) Haenke 


— 


1 


1 


Rorippa nasturtium- aquaticum (L.) Hayek 


Watercress 


1 


1 


Aubretia deltoidea (L.) DC. 


Aubretia 


1 


1 


Matthiola incana L. (R. Br.) 


Stock 


3 


2 


M. bicornis (Sibth. & Sm.) DC, 


Night-scented stock 


2 


2 


Malcolmia maritima (L.) R. Br. 


Virginia stock 


1/2 


1/2 


Hesperis matronalis L. 


Dame's violet 


2 


1 


Cheiranthus cherei L. 


Wallflower 


3 


2 


Camelina sativa (L.) Crantz 


Gold of pleasure 


1 


1 



11 

into grayish-white, irregular necrotic patches on the upper surface of the leaf while 
downy fungal growth appear on the under surface. In a severe attack, diseased leaves 
dry up and shrivel. On cotyledons, necrotic lesions are more pronounced in B. rapa, 
whereas on true leaves, lesions are conspicuous on B. juncea (Figs. 1, 2). 

Symptoms of mixed infection of downy mildew and white rust are common on 
leaves and inflorescence of B. rapa and B. juncea. On leaves, downy growth of the 
fungus appears in or around the white rust pustules (Fig. 3). On malformed 
inflorescences, sporulation of the downy mildew fungus is predominant in the form of 
white granular canidia and conidiophores (Fig. 4) (Saharan, 1992a). According to 
Butler (1918) owing to very frequent co-existence of white rust and downy mildew, it is 
not easy to separate their effects, but white rust produces the greatest deformities in 
the stem and flowers (Awasthi et al., 1995, 1997). Stem swellings may be limited often 
with abrupt bending of the stalk, or swelling may be several inches long. The axis of 
the inflorescence is equally susceptible to deformity. The leaves and flowers are not 
often swollen, except for the young ovary, which may be transformed to a twisted body 
about two or three inches in length. More often, the floral buds are atrophied with all 
the parts (i.e. sepals, petals, stamens, and pistal) being shrunken and almost colourless. 
If the attack is late, the buds/silique may be partly normal, partly deformed or 
atrophied, and a single bud may similarly be affected in part only. There is never any 
trace of the violet colour produced in downy mildew infections which often occur with 
white rust. 

Systemically mixed infected plants with P. parasitica and A. Candida have stunted 
and thickened growth of the whole plant which bears profuse sporulation of both 
pathogens. Hypertrophy of the affected cells, which is mainly attributed to infection 
with A. Candida, causes thickening of the stem and inflorescence. The hypertrophied 
tissues tend to attract infection by P. parasitica because their relative susceptibility to 
this pathogen is much higher than normal tissues (Awasthi et al., 1995, 1997). The pith 
of the stem has more hypertrophied tissue than the cortex. The affected inflorescence 
either bears no silique, or produces abnormal silique which are often curled without 
seeds. In the initial stages, an affected inflorescence does not show typical symptoms 
of infection such as the presence of oospores and downy growth on the surface. But at 
the later stages, conidial fungal growth of downy mildew and sporangiophores blisters 
of white rust occurs on the surface of affected tissue and formation of oospores takes 
place in the tissue as it dries. Necrotic lesions bearing downy growth of the fungus may 
also be observed on well developed silique (Awasthi et al., 1995, 1997; Kolte, 1985; 
Saharan, 1992a, b; Vasudeva, 1958). 

The internal changes due to infection by Peronospora differ from those caused by 
Albugo in many respects. With Peronospora infections, the palisade cells of the leaf are 
not changed. In the deeper layers of the cortex, endoderm and pericycle, new cell layers 




f 




J^^IylH 






IMrH 


Wfil&Kjttr 






h^m, 


f .^ 


K?>u^'"C»3S^B 




12 




■ fl 1 1 




W t?X ^M w* ? M* 








UmW* 




FSaSKL 


j 



Fig. 1 Irregular, necrotic lesions of downy mildew ( 1 a), and white growth of conidia and conidiophores of Pemiwspotv parasitica 

lb) on cotyledons of B.jimcea (Courtesy: Mehta, 1993). 
Fig. 2 Irregular, necrotic brown lesions of downy mildew on leaf lamina of B. juncea (Courtesy: Mehta, 1993). 
Fig. 3 Downy growth of Pervnospora parasitica in or around the white rust (Albugo Candida) pustules on the lower surface of 

B.jimcea leaf (Courtesy: Mehta; 1993). 
Fig. 4 White growth of conidia and conidiophores of Penmospora parasitica on Albugo Candida - induced malformed 

inflorescence of B. juncea (Courtesy: Mehta, 1993). 



13 



Table 3. Performance of the Brassica form of P. parasitica on cultivars of B. olerace 
(Reprinted from C.H. Dickinson and J.R. Greenhalgh. 1977. Host range and 
taxonomy of Peronospora of crucifers. Trans. Brit. Mycol. Soc. 69: 111-116, 
by permission of the authors and the publisher British Mycological Society) 







Mycelial 




Occurrence of pathogen 






development 




(% seedlings positive) 




Sporulation 


in cotyledon 
(%grid 
















Main 


Lateral 


Host Variety 


intensity* 


squares covered) 


Cotyledons Hypocotyl 


Root 


Root 


var. capitata L. (cabbage) 














cv. Red Drumhead 


+++ 


76 


100 


100 


90 


25 


cv. Savoy Drumhead 


+++ 


33 


100 


95 








cv. Flower of Spring 


++ 


31 


100 


100 








cv. Standby 


++ 


47 


100 


100 


25 





cv. Greyhound 


++ 


26 


100 


95 


5 





cv. Harbinger 


++ 


30 


100 


100 


45 





cv. Primo 


++ 


31 


100 


100 


35 





cv. January King 


+ 


11 


100 


60 








var. botrytis L. (cauliflower/broccoli) 












cv. Veitch's Self Protecting 


+++ 


63 


100 


100 


50 





cv. Roscoff Early 


+++ 


56 


100 


100 


5 





cv. Calabrese 


+++ 


53 


100 


100 


40 


5 


cv. All the Year Round 


++ 


50 


100 


100 


15 





cv. Veitch's Autumn Giant 


++ 


45 


100 


100 


25 


5 


cv. June 


++ 


23 


100 


100 








cv. Snowball 


++ 


50 


100 


100 








cv. Majestic 


++ 


24 


100 


95 








var. gemmifera Zenker, (brussel sprouts) 












cv. Masterman 


++ 


33 


100 


100 


10 





cv. British Allrounder 


++ 


37 


100 


100 


5 





cv. Cambridge No. 5 


++ 


51 


100 


100 


15 


5 


cv. Jade Cross 


++ 


18 


100 


100 








cv. Exhibition 


++ 


21 


100 


90 








cv. Cambridge No. 1 


++ 


34 


100 


75 








var. gongylodes L. (kohlrabi) 














cv. Green Vienna 


++ 


16 


100 


95 








cv. Purple 


++ 


20 


100 


100 








var. acephala D.C. (Borecole) 














cv. Tall Green 


++ 


42 


100 


95 


5 





cv. 1000-headed 


++ 


38 


100 


90 








subsp. oleracea (wild cabbage) 


+++ 


60 


100 


100 


60 


10 



* J. = 



+ = sparse sporulation; ++ = moderate sporulation; +++ = heavy sporulation 



14 



Table 4. Host species of P. parasitica (Channon, 1981; Verma, et al., 1994) 



Host 



Reference 



Arabidopsis spp. 

Armoracia rusticana 

Arabis spp. 

Aubretia spp. 

Brassica alba 

B. arvensis 

B. alboglabra 

B. chinensis 

B. juncea 

B. kaber 

B. hirta 

B. fructiculosa 

B. napus 

B. napus var. napobrassica 

B. nigra 

B. oleracea var. acephala 

B. oleracea var. botrytis 

B. oleracea var. capitata 

B. oleracea var. caularapa 

B. oleracea var. gemmifera 

B. oleracea var. gengyloides 

B. pekinensis 

B. campestris var. rapifera 

B. campestris var. yellow sarson 

B. campestris var. brown sarson 

B. campestris var. toria 

Chenopodium album 

B. tournefortii 
Barbarea 
Camelina sativa 
Cheiranthus allioni 

C. cheiril 

Capsella brusa-pasteris 
Cardamine impatiens 
C. rhomboidea 
Coronapus didymus 

C. squamatus 
Crambe maritima 
Dentaria spp. 

D. laciniata 
Descurainia spp. 
Draba spp. 

D. caroliniana 
Eruca sativa 
Hesperis spp. 



Horseradish 
Rockcress 
Aubretia 
White mustard 
Wild mustard 
Chinese kale 
Chinese cabbage 
Mustard 
White mustard 
White mustard 

Rape 

Swedes 

Black mustard 

Marrow stem kale 

Cauliflower 

Cabbage 

Brussels sprouts 
Kohlrabi 
Chinese cabbage 
Turnip 
Yellow sarson 
Brown sarson 
Toria 
Bathu 



Wallflower 



Crambe 



Taramira 



Koch & Slusarenko, 1990 

Moore, 1959 

Anonymous, 1960 

Moore, 1959 

Anonymous, 1960 a 

Anonymous, 1960a 

Johnston, 1963 

Hiura & Kanegae, 1934 

Gaumann, 1926 

Anonymous, 1960a 

Anonymous, 1960a 

Gaumann, 1926 

Moore, 1959 

Moore, 1959 

Gaumann, 1926 

Moore, 1959 

Moore, 1959, Ramsey, 1935 

Moore, 1959, Ramsey, 1935 

Moore, 1959, Ramsey, 1935 

Moore, 1959, Thung, 1926 

Johnston, 1963 

Chang et al., 1963, Ocfemia, 1925 

Moore, 1959 

Saharan, 1992a 

Saharan, 1992a 

Saharan, 1992a 

Saharan, 1996 

Gaumann, 1926 

Anonymous, 1960a 

Darpoux, 1945 

De Bruyn, 1935b 

Moore, 1959, Wiese, 1927 

Farlow, 1883 

Sharma & Munjal, 1977 

Farlow, 1883 

Langdon, 1948 

Dias & Da Camara, 1953 

Moore, 1959 

Anonymous, 1960a 

Farlow, 1883 

Anonymous, 1960a 

Anonymous, 1960a 

Farlow, 1884 

Gaumann, 1926 

Anonymous, 1960a 



15 



Erysimum cheiranthoides 


Warmseed mustard 


Anonymous, 1960a 


Iberis amara 


Candytuft 


Anonymous, 1960a 


Lepidium sativum 


Garden cress 


Anonymous, 1960a 


L. intermedium 




Swingle, 1890 


L. graminifolium 




Nicolas & Aggery, 1940 


L. virginicum 




Farlow, 1883 


Lobularia spp. 


Koniga 


Anonymous, 1960a 


Malcolmia africana 




Thind, 1942 


Matthiola incana 


Stock 


Moore, 1959; Wiese, 1927 


Nasturtium officinale 


Watercress 


Moore, 1959 


Raphanus sativus 


Radish 


Moore, 1959 


R. raphanistrum 


Wild radish 


Gaumann, 1926 


Radicula nasturtium-aquaticum 




Davis, 1929 


Rorippa 




Anonymous, 1960a 


Sinapis alba 


White mustard 


Gaumann, 1926 


S. arvensis 


Wild mustard 


Gaumann, 1926 


Sisymbrium officinale 


Hedge mustard 


Anonymous, 1960a 


S. irio 


Tumbling weed 


Thind, 1942 



may be formed by normal cell division. Only the cells around the vascular bundles 
become enlarged and thin-walled; the rest of the interfascicular sclerenchyma remains 
unaltered. There is no interfascicular cambium in these host plants. The cambium of 
the vascular bundles remains active whereas the xylem and phloem vessels become 
enlarged and separated by radial bands of parenchyma when hyphae have penetrated. 
There are no accessory bundles. In general, the effect on the cells seems to be more 
destructive than in Albugo, the chlorophyll content is diminished and the cell contents 
more rapidly used up. There is no tendency for chlorophyll accumulation in unusual 
places as with white rust. In general, the effect of downy mildew on the cell seems to 
be more destructive than white rust (Butler, 1918). 

The protoplast of epidermal cells respond differently to haustorial development 
than protoplast of the mesophyll cells (Chau, 1970). The epidermal cells result in a 
severe disruption of the protoplast. The central vacuoles contract and probably undergo 
fragmentation, the plasmalemma is broken down or detached from the wall and 
numerous vesicles are formed from it. The cytoplasm is either dislocated and 
aggregated into a vacuolated blob or completely dispersed to the extent that its identity 
cannot be discerned. Consequently, haustoria in epidermal cells are not, in most cases, 
surrounded by a clearly defined layer of host cytoplasm. Haustoria formation in a 
mesophyll cell cause less disruption. The host cytoplasm is merely invaginated by the 
invading haustorium, while the tonoplast and plasmalemma apparently remain intact. 



ii) Brassica vegetables: Plants can be infected at any time (Sherf and Macnab, 
1986). In seed beds, the cotyledons and first leaves are invaded. The adaxial surface 



16 

of the leaf bears small, pale-yellow, angular spots, which may grow together to form 
irregular brown patches. On the abaxial surface the corresponding areas are covered 
with a light-gray fungus formed by multi-branched conidiophores bearing conidia. 
Young leaves and cotyledons may drop off as they yellow. Older leaves usually persist, 
and affected areas enlarge becoming papery and tan coloured. Severe infection may 
cause the death of the whole leaves. Minute necrotic flecks covering the leaf surface 
may often form resembling peppery leaf spot caused by bacteria. 

When the fungus enters the stalk at the leaf base of an old head of cabbage, a 
grayish-black discolouration of the stalk occurs (Ramsay and Smith, 1961). In some 
storage lots of cabbage this discolouration has been found extending up through the 
stalk to the innermost bud leaves. On cabbage heads, the pathogen may cause 
numerous sunken black spots, varying in size from minute dots to an inch or more in 
diameter (Sherf and Macnab, 1986). A similar blackening occurs on cauliflower curds. 
The infection is evident as brown to black streaks in the vascular system of the upper 
portion of the main stalk and branches leading to the florets. The fleshy roots of turnips 
and radishes have an internal irregular region of discolouration extending from the root 
crown downward or beginning on the side at soil level. The flesh is brown to black or 
shows net necrosis. In advanced stages the skin can be roughened by minute cracks, 
and the root can split open (Sherf and Macnab, 1986). 

According to Butler (1918) the fungus is visible as a thin, grayish-white, downy 
growth, occurring in scattered patches on the under surfaces of the leaves in cabbage, 
cauliflower and turnip, and on the leaves, stem and inflorescence in radishes. The 
upper surface of the leaf is marked by white spots corresponding to the downy growth 
below. In severe attacks, the spots may be so crowded that the leaf dries up, shrivels, 
and tears easily. In seedlings, the whole under surface may be evenly covered, and total 
infection of the young inflorescence is also found. Occasionally the roots of radish and 
Swedish turnip are attacked in Europe. The tissues blacken and rot near the surface, 
oospores occur within the tissues and conidiophores form if exposed to the air. 

iii) Broccoli: Downy mildew appears first on the lower leaves of broccoli plants 
(Natti et al., 1956). Leaf infection may occur soon after the plants are set in the field 
or may take place later in the season. Older leaves appear to be more susceptible than 
newly developed leaves. When the surface of the foliage is wet, the downy white 
mycelium of the fungus is readily observed on the under surface of the leaves. The first 
symptoms of leaf infection are small water-soaked spots surrounded by a halo of light 
green tissue on the under surface of the leaf. Under conditions favourable for 
development of infection, the spots enlarge to form indefinite yellow areas. Later, the 
tissues within these infected areas collapse and become light brown and parchment-like. 
The mildew lesions vary in size and shape. The largest lesions usually are bounded by 
leaf veins. The initial spots of infection may also remain localized. The tissues of the 



17 

spot collapse to form a small brown lesion. Systemic infections are usually confined to 
the upper portion of the main stalk and to the branches leading to the florets of the 
head. Infected tissues develop brown to black netted lesions, and in others as long 
strands of discoloured tissues. In some plants systemic invasion of the head can be 
detected by diffuse blue to purple areas on the stalk and branches of the head. 

iv) Wallflower (Cheiranthus) : On the diseased plants the upper surface of the 
leaves show pale yellowish patches, while the corresponding parts of the undersurface 
are covered with a grayish or white fungal growth (Gram and Weber, 1952). The 
infected stems and flowers are swollen and often twisted. Diseased flower buds do not 
develop. 

v) Stock (Matthiola): The disease is more common on young plants before they 
are transplanted but may also appear later, especially on crop grown indoors (Gram and 
Weber, 1952). On the upper surface of the leaves there are pale spots, while on the 
corresponding parts of the undersurface is a whitish layer of the fungus. Stalks and 
flower heads may also be attacked. The diseased parts show various kinds of distortion. 
According to Jafar (1963) the disease appears as light green areas on the upper surface 
of leaves. The corresponding under surface is chlorotic with white growth of fungal 
conidia and conidiophores. Infected areas turn yellow, and become necrotic leading to 
premature leaf fall. The flowers of infected plants frequently fail to open and often die. 
Fructification appear on cotyledons and seedlings may be killed. 

e. Disease assessment 

Different scales have been used for classifying leaf infection by downy mildew 
pathogen. Natti et al., (1967) and Sadowski (1987) used scales ranging from 0-5, where: 

= no symptoms 

1 = spots, necrotic flecks or streaks, but no sporulation 

2 = spots, necrotic flecks or streaks, with sparse sporulation confined to necrotic 

tissue 

3 = systemic infection and sporulation in 

4 = systemic infection and sporulation in increasing degree 

5 = systemic infection and sporulation in increasing degree 

Plants with ratings to 2 are considered resistant. Ebrahimi et al. (1976) has 
rated downy mildew resistance in B. juncea lines on a scale of 1 to 5 where, 1 = 
indicates no sporulation, 2 = very sparse sporulation, and 5 = heavy sporulation. A 
similar scoring scale of 1 to 4 with slight modification has been used by Dickinson and 
Greenhalgh (1977). Use of a - 9 scale has been suggested by several workers (Knight 
and Furber, 1980; Nashaat and Rawlinson, 1994; Saharan, 1992b; Williams, 1985). It 



18 

is described as follows: 

= no symptoms or signs of P. parasitica 

1 = very minute to larger scattered necrotic flecks under the inoculum drop, no 

or small amounts of necrosis on the lower cotyledon surface, no sporulation 
3 = very sparse sporulation, one to a few conidiophores on the upper or lower 

surfaces, necrotic flecking often present, tissue necrosis present 
5 = sparse scattered sporulation on either or both cotyledon surfaces, tissue 

necrosis 
7 = abundant to heavy sporulation mainly on lower surfaces, light to scattered 

sporulation on upper surfaces; tissue necrosis and chlorosis may be present 
9 = abundant sporulation; leaf or cotyledon collapsed. 

A disease index (DI) was calculated using the formula: 

9 
DI= I(ixj)/n 

i=0 
where n = total plants, i = infection phenotype class, and j = number of plants per class. 
Genotypes are categorized as resistant (0 to 1), partially resistant (3 to 5) and 
susceptible (7 to 9). 

Kruger (1991) suggested use of 1-9 scale in the form of diagrams to estimate 
disease on leaves of oilseed rape (Fig. 5); scores 3, 5, 7, and 9 respectively represent 7, 
27, 65, and 100 percent of the leaf area infected. If larger leaves are concerned, those 
in Fig. 5 should be enlarged by 2 to 5 times to get a comparable shape and size to the 
leaves found in the field. 

Brophy and Laing (1992) assessed disease severity using an image analyser to 
determine logarithmic rating scales of percentage leaf area infected for both cotyledons 
and primary leaves of cabbage. They found that the maximum area infected can be 
100% in cotyledons but in primary leaves it rarely exceeds 25%. In order to integrate 
the two components, transformation of the data is necessary. Percentage disease 
severity (PDS), expressed as a function of cotyledon and primary leaf infection is 
calculated using the formula PDS = (C + x P)/2, where C is the percentage cotyledon 
area infected with a maximum value of 100%; x is the inverse of the maximum 
measured percentage primary leaf area infected, and P is the percentage primary leaf 
area infected of treated plants. 



3. THE PATHOGEN 

Downy mildew of crucifers is caused by an obligate pathogen, Peronospora 



19 




Sco«t J <«fncUD till 71 ) 



Fig. 5. Disease assessment (1 to 9) on leaves of oilseed rape (Reprinted from W. Kriiger. 1990. A review on assessment of 
diseases in oilseed rape- comparison of various methods. IOBC/WPRS Bulletin, Number 14: 91-1 1 1, by permission of the 

author and the publisher). 



20 

parasitica (Pers. ex. Fr.) Fr., Sum. Veg. Scand. 493, 1849. Extensive synonymy is given . 
by Yerkes and Shaw (1959). Sometimes it is referred to as Peronospora brassicae. 

a. Taxonomy and morphology 

The earliest reference of downy mildew on crucifers is by Persoon (1796) who 
ascribed the cause of the disease on Thlaspeos bursa-pastoris (Capsella bursa-pastoris) 
to the fungus Botrytis parasitica Pers. In 1849, Fries (Gaumann, 1918) transferred the 
fungus to the genus Peronospora which had been established in 1837 by Corda in his 
description of Peronospora ramicis (Corda, 1837). At that time all isolates obtained 
from cruciferous hosts were ascribed to P. parasitica (Pers. ex. Fr.) Fr. However, 
Gaumann (1918) named isolates of Peronospora affecting plants of Brassica species as 
P. brassicae Gaum. He considered that the various isolates obtained from different 
hosts should be classified as separate entities, and on this basis recognized 52 species 
of Peronospora. His conclusions were based largely upon conidial dimensions and the 
results of cross-inoculation tests. The value of conidial dimension as a taxonomic 
criterion has since been questioned because size may vary according to environmental 
conditions (Thung, 1926a). Yerkes and Shaw (1959) reported remarkable morphological 
similarity of Peronospora species which attack crucifers. Measurements of conidia 
(Tables 5, 6), an inability to associate conidiophores types with particular host genera, 
and the uniformity of oospores led Yerkes and Shaw (1959) to conclude that there is no 
reliable morphological basis to distinguish different species of Peronospora affecting the 
crucifers. Following an extensive biometric study, over 80 species names were reduced 
to one synonym and now a single species, i.e., P. parasitica, has been recognized on 
cruciferae hosts (Dickinson and Greenhalgh, 1977; Haura and Kanegae, 1934; 
Waterhouse, 1973; Yerkes and Shaw, 1959). However, in view of the apparent 
differences in the antheridial structure in the isolates of Peronospora on Capsella bursa- 
pastoris (Wager, 1889) and on B. oleracea (McMeekin, 1960) the merits of some separate 
speciation must not be ruled out. The phylogeny of the Peronosporales (Fig. 6) has been 
shown by Shaw (1981). 

b. Reproduction and reproductive structures 

The general morphology and infection cycle of P. parasitica is similar to that of 
other members of the family Peronosporaceae. 

i) Asexual phase: Mycelium and haustoria : The mycelium is hyaline and 
coenocytic. It grows intercellularly in the host tissues and produces haustoria to 
penetrate the host cells. The haustoria are large, lobed, elongated or club shaped 
(Butler, 1918; Fraymouth, 1956; Holliday, 1980). They branch extensively and can 
nearly fill the entire cell. In the leaf of Japanese radish, the mycelia turn and twist 
irregularly in the intercellular spaces of the spongy parenchyma, and usually develop 



21 



ON GRAMINEAE 



ON OTHER ANGIOSPERMS 



Peronosclerospora 



Peronospora 

ii 



Bremia 



Bremia 



Sclerospora 




Peronospora 
leptosperma 



Pseudoperonospora 

ii 



Bosidiophoro 



Plosmopara 3 
oplismeni 

Plosmopora 3 
penmseti 



Sclerophlhora' 




Rhysofheca 
(Plasmopora 
Sensu la to) 



Plasmopara 
cephalophora 





Bremiello 

Albugo 



Peronophylhora 



Phylophthoro 

ii 



Pylhium 



Fig. 6. Phylogeny of the Peronosporales (Reprinted from C.G. Shaw. 1981. Taxonomy and evolution. ]N. The downy mildews, 
D.M. Spencer (editor) Chapter 2: 17-29, by permission of the author and the publisher Academic Press Limited, London). 



22 



Table 5. Measurements of Peronospora conidia on Crucifers (Reprinted from 
W.D. Yerkes and C.G. Shaw. 1959. Taxonomy of the Peronospora 
species on cruciferae and chenopodiaceae. Phytopathology 49:499-507, 
by permission of the authors and the publisher American 
Phytopathological Society) 





r 










Quotient 










Mo. spores 






Length (u) 


length/width) 
Means 




Wi 


measured 




Min. 


Means 
Grand 


Max 


Range 


Min. 


Means 
Grand 


Max. 


Range 


+ no. 




Min. 


Grand 


Max. 


collections 


Sisymbrium altis- 


























simum L. 




(15.66) 




11.9-19.9 




(1.32) 






(11.89) 




9.6-15.3 


100 (1) 


Arabis hirsuta 




(16.12) 




11.0-23.0 




(1.16) 






(13.92) 




8.0-20.0 


1,000 (1) 


Arabis laevigata 


























(Muhl.) Poir. 




(16.89) 




10.3-16.1 




(1.24) 






(13.63) 




12.6-19.9 


100 (1) 


Arabis hirsuta (L.) Scop. 


16.99 


(17.62) 


18.11 


13.4-22.6 


1.23 


(1.30) 


1.37 


13.02 


(13.54) 


14.48 


11.1-19.2 


220 (3) 


Cardamine bulbosa 


























(Schreb.) BSP. 




(17.72) 




13.4-23.0 




(1.25) 






(14.22) 




11.5-16.1 


100 (1) 


Draba caroliniana 




(17.64) 




11.0-24.0 




(1.12) 






(15.81) 




9.0-21.0 


1,000 (1) 


Rorippa palustris (L.) 


























Bess. 




(18.75) 




15.3-24.9 




(1.31) 






(14.27) 




11.5-18.0 


100 (1) 


Cardamine parviflora L. 




(18.90) 




14.2-23.0 




(1.28) 






(14.79) 




11.5-17.2 


100 (1) 


Nasturtium officinale 


























R. Br. 


18.32 


(19.50) 


20.29 


14.6-26.8 


1.17 


(1.21) 


1.25 


15.33 


(16.19) 


17.37 


12.3-21.1 


140 (3) 


Cardamine penn- 


























sylvanica Muhl. 




(19.62) 




15.3-23.8 




(1.35) 






(14.56) 




12.6-16.9 


100 (1) 


Brassica nigra (L.) Koch 




(19.77) 




15.7-23.7 




(1.16) 






(17.00) 




14.5-21.4 


100 (1) 


Brassica arvensis (L.) 


























Ktze. 




(19.97) 




15.3-27.2 




(1.25) 






(16.04) 




11.5-20.7 


100 (1) 


Raphanus sativus 




(20.01) 




14.0-26.0 




(1.10) 






(18.17) 




14.0-22.0 


1,000 (1) 


Draba caroliniana Walt. 


19.53 


(20.19) 


21.37 


14.6-26.4 


1.21 


(1.22) 


1.25 


15.68 


(16.58) 


17.66 


11.9-20.7 


220 (3) 


Nasturtium officinale 




(20.32) 




16.0-27.2 




(1.19) 






(17.07) 




12.8-20.8 


101 (1) 


Dentaria laciniata Muhl. 


20.23 


(21.17) 


22.87 


15.3-27.6 


1.26 


(1.29) 


1.31 


15.69 


(16.45) 


17.41 


11.5-21.8 


140 (3) 


Capsella bursa-pastoris 


20.87 


(21.39) 


21.90 


12.0-35.0 


1.15 


(1.18) 


1.20 


18.21 


(18.26) 


18.30 


11.0-24.0 


1,200 (2) 


Capsella bursa-pastoris 


























(L.) Medic. 


20.79 


(23.10) 


28.40 


14.6-36.8 


1.26 


(1.35) 


1.44 


15.86 


(17.14) 


19.70 


11.9-23.8 


1,200(13) 


Raphanus sativus L. 




(23.54) 




18.0-29.5 




(1.26) 






(18.72) 




14.2-22.6 


100 (1) 


Sisymbrium canescens 




(26.38) 




17.0-32.0 




(1.36) 






(13.25) 




12.0-24.0 


1,000 (1) 


Sisymbrium canescens 


























Nutt. 




(27.13) 




19.9-33.3 




(1.77) 






(15.33) 




11.9-18.4 


100 (1) 


Lepidium virginicum 




(27.52) 




19.0-37.0 




(1.50) 






(18.35) 




14.0-24.0 


1,000 (1) 


Lepidium apetalum 


























Willd. 


28.94 


(29.19) 


29.38 


19.1-38.3 


1.46 


(1.47) 


1.48 


19.44 


(19.79) 


20.14 


15.3-25.3 


120 (2) 


Lepidium virginicum L. 




(29.46) 




23.0-38.3 




(1.50) 






(19.68) 




15.7-26.8 


100 (1) 



23 



Table 6. Measurements of Peronospora conidia on Chenopodiaceae (Reprinted from 
W.D. Yerkes and C.G. Shaw. 1959. Taxonomy of the Peronospora species on 
cruciferae and chenopodiaceae. Phytopathology 49:499-507, by permission 
of the authors and the publisher American Phytopathological Society) 













Quotient 








No. spores 






Length (u) 


lenerth/width) 
Means 




Width (u) measured 




Min. 


Means 
Grand 


Range 
Max. 


Min. 


Means 
Grand i 


Range 
Max. < 


+ no. 




Min. 


Grand 


Max. 


:ollections 


Spinacia oleracea 




(24.30) 


17.6-32.0 




(1.22) 






(19.90) 


12.8-25.6 


500 (1) 


Beta vulgaris L. 


21.54 


(24.81) 


27.69 17.5-32.5 


1.25 


(1.30) 


1.36 


17.13 


(18.94) 


20.88 13.7-23.7 


375 (15) 


Chenopodium murale 




(24.83) 


16.0-32.0 




(1.14) 






(21.63) 


12.8-28.8 


500 (1) 


Chenopodium hybridum 




(24.98) 


17.6-32.0 




(1.46) 






(17.11) 


8.0-24.0 


500 (1) 


Chenopodium bonus- 






















henricus 




(26.01) 


17.0-34.0 




(1.13) 






(22.86) 


16.0-31.0 


500 (1) 


Chenopodium sp. 


24.88 


(26.55) 


28.72 20.0-35.0 


1.30 


(1.34) 


1.40 


19.09 


(20.09) 


20.46 16.2-26.2 


100 (4) 


Chenopodium lepto- 






















phyllum Nutt. 




(26.68) 


20.0-33.7 




(1.38) 






(19.28) 


15.0-23.7 


25(1) 


Spinacia oleracea L. 


24.43 


(26.92) 


28.99 20.0-37.5 


1.26 


(1.37) 


1.47 


18.08 


(19.61) 


20.74 15.0-25.0 


750 (30) 


Chenopodium bonus- 






















henricus L. 


26.23 


(26.95) 


27.67 22.5-33.7 


1.28 


(1.30) 


1.33 


20.49 


(20.70) 


21.32 16.2-23.7 


125 (5) 


Chenopodium hybrid- 






















ium L. 


23.94 


(27.53) 


33.44 16.9-40.2 


1.30 


(1.38) 


1.50 


17.11 


(19.88) 


22.23 15.3-25.7 


150 (6) 


Chenopodium giganto- 






















spermum Aeller 




(28.58) 


26.2-36.5 




(1.46) 






(19.57) 


17.5-21.2 


25(1) 


Chenopodium murale L. 




(29.09) 


25.0-35.0 




(1.31) 






(22.17) 


16.2-26.2 


25(1) 


Chenopodium album 




(29.10) 


17.2-40.2 




(1.53) 






(19.04) 


12.2-27.2 


500 (1) 


Chenopodium album L. 


23.83 


(29.65) 


33.53 20.0-40.0 


1.31 


(1.41) 


1.58 


17.77 


(20.98) 


23.17 14.6-27.5 


825 (33) 



only one haustorium for each host cell (Ohguchi and Asada, 1990). However, in root 
tissues where parenchyma cells are large and much closer together, the mycelia are 
smooth and one to several haustoria are formed in the infected host cell. Mycelial 
growth patterns in petioles and hypocotyls are similar to those in root tissue. Prior to 
haustorium formation a leaf like structure is formed from the intercellular mycelium 
in the narrow spaces between root parenchyma cells. It is flat, 6 /mi thick and covers 
the surface of host cell. The leaf like structure forms various type of haustoria, ranging 
from 0-25 number in one cell. 



In turnip and radish roots, the haustoria are initially spherical to pyriform, but 
later become cylindrical or clavate, and often dichotomously or trichotomously branched 
(Chu, 1935). In cabbage, some haustoria are large irregular vesicles while others are 
bilobed and regular in shape. In cauliflower, they are single, globose and uniform in size. 



24 

Variations in shape and size of haustoria of P. parasitica occur in hosts other than 
Brassica spp., such as Matthiola incana, Cheranthus cheiri, Capsella bursa-pastoris, 
Diplotaxis muralis and Rhynchosynapis manensis (Fraymouth, 1956). Penetration of 
the haustorial branch occurs through a hole 1-2 fx diameter in the cell wall which may 
form a collar-like structure round the base of the primordial haustorium. As the 
haustorium enlarges, invagination of host plasmalemma occurs and a sheath, possibly 
of callose, forms round the intrusive organ (Fraymouth, 1956). Moderately high 
temperatures of 20-24°C favour the most rapid development of the haustoria (Felton 
and Walker, 1946). 

r 

Conidiophores and conidia : After vegetative growth of the mycelium, erect 
conidiophores singly or in groups emerge vertically through stomata on the abaxial 
surface of the host leaves during a period of darkness. The conidiophores are hyaline 
and measure 200-300 jj,. Condiophores are uniform with a flattened base and stout 
main axis. At 8°C, the rate of elongation reaches 100-200 /j. h" 1 and the whole process 
from emergence to spore formation takes approximately 4-6 h (Davison, 1968b). They 
are dichotomously branched, 6-8 times, tips bifurcate, branching acute and slightly 
thickened above each fork. The terminal branches are long, slender, pointed and end 
in a single conidium. The sterigmata are slender and acutely pointed (Butler, 1918; 
Holliday, 1980; Channon, 1981). 

The conidia are hyaline, broadly elliptic, or nearly globose, measure 24-27 x 15-20 
\x and are delimited from sterigmata by cross-walls at maturity. A single conidium is 
borne at the tip of each branch and is deciduous (Butler, 1918; Holliday, 1980). 
Detachment of conidia is possibly caused by hygroscopic twisting of the conidiophores 
which in turn is related to fluctuations in humidity (Pinckard, 1942). Conidia 
germinate in free water by a lateral germ-tube, not by zoospores. Infection occurs both 
by direct penetration of the epidermis and through stomata (Butler, 1918). In 
cauliflower leaves conidia form appressoria in the junction areas between the anticlinal 
walls of adjoining epidermal cells (Preece et al., 1967). 

ii) Sexual phase: Sexual organs, gametogenesis. fertilization and oospore 
formation : During sexual reproduction P. parasitica forms spherical oogonia and 
paragynous antheridia. Oogonia are pale yellow, irregularly round, and swollen into 
crestlike folds (Butler, 1918; Holliday, 1980). Antheridia are tendril-like and are 
produced on separate hyphae. Wager (1900) observed that the protoplasm of the 
oogonium becomes differentiated into a central vacuolated ooplasm and a peripheral 
multinucleate granular periplasm. A receptive thin-walled papilla forms on the 
oogonium at the point of contact with the antheridium. A fertilizing tube grows from 
the antheridium through the receptive papilla towards a "central body" in the ooplasm, 



25 

to discharge a single "male" nucleus. Meanwhile, a single "female" nucleus detaches 
itself from the periplasm and also migrates towards the central body. The two nuclei 
fuse and initiate the uninucleate oospore. During ripening of the oospore the periplasm 
is deposited on its wall as an exosporial layer. The oospores are formed in the host 
tissues at late stage of sporulation. They have also been found in the cavity of the ovary 
on hyphae emerging between the cells of the inner epidermis of the carpels. The 
oospore lies inside, almost filling the cavity. The mature oospore is thick-walled, 
yellow-brown and globose or spherical, and measures 30-40 /u in diameter (Butler, 1918; 
Holliday, 1980). Oospore formation is favoured by conditions which induce senescence 
of the host tissues such as a deficiency of N, P, or K (McMeekin, 1960). Germination of 
oospores is by a germ-tube (Butler, 1918). 

c. Electron microscopy and ultra structures 

Electron microscopy in association with physiological, biochemical and genetical 
studies have provided information which helps in understanding the complex host- 
parasite relationship of this disease. 

i) Host penetration: Penetration and haustorial formation in epidermal cells 
begins 6h after inoculation (Fig. 7B, C) (Chou, 1970). Appressoria, which look like 
swollen discs 7-10 /u, across form at the junction of epidermal cells (Fig. 7B). At this 
stage, the appressoria and haustoria appear densely granulated, as the spores empty 
their contents during the process of germination and infection. The penetrating hyphae 
lay in between the anticlinal cell walls of the two epidermal cells (Fig. 7C) along with 
the formation of one or two haustoria, reaching to the adjacent mesophyll cells. After 
45h, intercellular hyphae ramify through more cells and reach the opposite epidermis. 
At this stage, the haustoria appear broad and conspicuous, reaching 20^ in length. The 
intercellular hyphae are about 7/u, across. Sometimes a sheath can be observed 
enveloping a fully grown haustorium (Fig. 7D). The earliest detectable stage of 
penetration is the formation of penetration hyphae which are as long as the vertical 
depth of the entire epidermal cell. The thick wall of the appressorium, continuous with 
the wall of the penetration hypha (Fig. 7A, 9A), has only a thin peripheral layer of 
cytoplasm. The cell contents migrate into the newly formed penetration hypha. In 
some cases, the appressorium can be seen to be embedded in an electron-dense, 
vacuolate material (Fig. 9B) appearing to be a mucilaginous sheath. This sheath is 
bound by an outer membrane which adheres to the cuticle of the host epidermis with 
the exception of in the penetration region where it is slightly separated (Fig. 9B). The 
penetrating hyphae wedge into the middle lamella between the anticlinal walls of two 
epidermis cells. The hole in the wall through which the fungus penetrates is 4-5/U 
across. After entering the host, the hypha expands to a diameter of 7-8/u.. There is no 
clearing zone or dissolution of wall material in the immediate vicinity of the penetrating 
hypha. The penetrating hypha is always seen to be embedded in a moderately electron - 




26 

Fig. 7. (A) Electron micro- 
graph of T.S. of epidermal 
cells of cabbage cotyledon at 
6-h after inoculation 
showing appressorium (ap) 
and penetrating hypha of 
Peronospora parasitica in 
between the anticlinal walls (j) 
of host epidermal cells. In one 
of the cells a haustorium was 
formed but the section only 
shows part of sheath(s). The 
penetration was cut obliquely 
and part of the hyphal wall 
(arrow pointed) is shown, x 
8200; (B) Photomicrograph of 
whole mount of a cleared 
cabbage cotyledon at 6-h after 
inoculation showing 

appressorium (ap) formation 
predominantly at the junction 
lineof epidermal cells. x313; 

(C) Photomicrograph C.T.S. 
of cabbage cotyledon at 6-h 
after inoculation showing 
penetration as in A. x 500; 

(D) Photomicrograph of whole 
mount of a cleared cotyledon 
showing intercellular hypha 
and haustorium completely 
ensheathed. x 840 (Reprinted 
from C.K. Chou. 1970. An 
electron microscope study of 
host penetration and early 
stages of haustorium 
formation of Peronospora 
parasitica (Fr.) Tul. on 
cabbage cotyledons. Ann. 
Bot. 34: 189-204, by 
permission of the author and 
the publisher Academic Press 
Limited, London). 



Abbreviations for Figures 7-17: 



ap = appressorium; apw - appressorium wall; ch = chloroplast; cu = cuticle; cy = cytoplasm; d = dense granules; Ep = host 
epidermal cell; f = hypha; fw = hyphal wall; h = host; ha = haustorium; hai = haustorium initial; hap = haustorium 
plasmalemma; haw = hautorium wall; hp = host plasmalemma; ht = host tonoplast; hw = host wall; j = anticlinal wall or 
junction line of host epidermal cells; lo= lomasome; m = membrane; ma = matrix of dense layer; me = mesophyll cell; mi 
= mitochondria; mu = mucilaginous sheath or substance; n = nucleus; ne = neck of haustorium; pef = penetrating hypha; s 
= sheath; smx = sheath matrix; v = vacuole; wl = cuticular layer; w2 = wall proper; z = dense zone: zl = outer dense zone; 
z2 = inner dense zone; 



27 




Fig. 8. (A) Electron micrograph of T.S. of epidermal cells of cabbage cotyledons at 8i-h after inoculation showing intercellular 
hyphae at various stages of penetration to the outside of host epidermis. Arrow points at the spearhead-like thickening of 
hyphal tip. x 5400; (B) Electron micrograph of part of outgrowing hypha in between two host epidermal cells showing die- 
back of hyphal tip and walling-off (arrow pointed) of apparently intact cytoplasm, x 18000 (Reprinted from C.K. Chou. 
Z1970. An electron microscope study of host penetration and early stages of haustorium formation of Peronospora 
parasitica (Fr.) Tul. on cabbage cotyledons. Ann. Bot. 34: 189-204, by permission of the author and the publisher 
Academic Press Limited, London). 



28 




Fig. 9. (A) Electron micrograph of T.S. of epidermal 
cells of cabbage cotyledon at 6-h after 
inoculation. The penetration region was cut 
medianly through showing the appressorium is 
almost empty with cytoplasm migrating into the 
penetrating hypha. x 14400; (B) Electron 
micrograph of T.S. of appressorium and part of 
host epidermal cells showing the mucilaginous 
sheath of the appressorium. Membranous 
boundary of the mucilaginous sheath is shown by 
arrow, x 6000: (C) Electron micrograph of 
section of a haustorium initial in host epidermal 
cell. Pan of the neck of a fully grown 
haustorium is shown by its side. Note the hyphal 
wall is continuous with wall of the intercellular hypha at this stage, x 13800: (D) Electron 
micrograph of a section of intercellular hypha and host epidermal cell showing pan of host wall in contact with hypha 
is swollen and partially eroded (arrow), x 17700 (Reprinted from C.K. Chou. 1970. An electron microscope study of 
host penetration and early stages of haustorium formation of Peronospora parasitica (Ft.) Tul. on cabbage cotyledons. 
Ann. Bot. 34: 1 89-204. by permission of the author and the publisher Academic Press Limited. London). 



29 

dense matrix of the middle lamella (Fig 7A, 9A). The cuticle breaks and fits closely 
around the penetrating hypha. No sign of swelling or change in electron density of the 
cuticle can be detected in the immediate vicinity of the penetration zone (Chou, 1970). 

After 80h, hyphal growth develops conidiophores which may be seen coming out 
from the epidermal cells (Chou, 1970). The intercellular hyphae appear to aggregate 
beneath the epidermis and grow either through stomata or in between two epidermal 
cells to the outside of the host tissue (Fig. 8A). An electron-dense spearhead-like 
thickening of the hyphal tip is observed to wedge in between two guard cells. This 
thickening may give rigidity to the hyphal tip for penetration. The hyphae are 
cemented to each other and also to the host cell walls by an amorphous, moderately 
electron-dense material, presumably of a mucilaginous nature (Fig. 8A, B). Hyphae 
penetrating through the junction of epidermal cells invariably show a die-back of the 
tip (Fig. 8A). A new wall is laid down round the remaining living cytoplasm, while a 
new growing tip is organised to carry on further growth (Fig. 8B). Large numbers of 
lomasomes appear around the newly formed walls and numerous dense vesicles 
approximately 500-1000A in diameter are concentrated in the walled-off cytoplasm. 

ii) Haustorium development: Host penetration by haustoria of the 
Peronosporales is usually by boring a narrow canal at the point of contact between the 
hypha and the host cell wall (Fraymouth, 1956). However, according to Chou (1970), 
it is not possible to find the stage at which the walls of both host and pathogen are 
perforated prior to haustorial initiation. Localized swelling of the host wall (3X 
original) is observed in the area of hyphal contact. The swollen area is about 1.5-2// 
long, and shows a clearer fibrillar structure, with a partially eroded area (Fig. 9D). The 
dimension of the swollen region coincides closely with the size of the hole in the host 
wall made by the haustorium. These observations strongly suggest that the breach of 
host wall during haustorium initiation is achieved at least partly by chemical means. 
A dome-shaped protuberance, about 1// in diameter is formed by the bulging of the wall 
of intercellular hypha into the lumen of host cell (Figs. 9C, 10A). The host wall is 
perforated at this stage and the wall of the haustorium primordium is continuous with 
that of the intercellular hypha (Fig. 9C). The haustorium initial is completely enclosed 
in a mound-like sheath quite distinct from the host wall in structure as well as density. 
The perforation made by the invading haustorium measures l-2,a across. The perforated 
host wall in most cases remains smooth, but a slight infolding of the wall to form a short 
collar-like structure is sometimes observed. The external part of the wall of the 
haustorial initial consists of a very electron-dense layer varying in thickness from 0.1- 
0.2// and exhibiting an undulating surface bounded externally by a thickened 
membrane (Figs. 9C, 10A) which is presumably the invaginated plasmalemma of the 
host. The primordial haustoria is filled with homogeneous ground-plasm packed with 
ribosomes. Lomasomes are the only organelles present at this stage. The growth and 
differentiation of the primordial haustorium is in the form of an elongated neck and 




30 



Fig. 10. (A) Electron micrograph of a section of a haustorium initial in host mesophyll cell (section slightly oblique to the 
penetration zone), x 16500; (B) Electron micrograph of a section of a very young haustorium in host epidermal cell 
showing breakdown of host cytoplasm into large number of vesicles, x 18000 (Reprinted from C.K. Chou. 1970. An 
electron microscope study of host penetration and early stages of haustorium formation of Peronospora parasitica (Fr.) 
Tul. on cabbage cotyledons. Ann. Bot. 34: 1 89-204, by permission of the author and the publisher Academic Press Limited, 
London). 



31 

expanding head (Fig. 10B). The sheath seems to burst apart, remaining as a collar-like 
structure around the neck region (Fig. 10B). 

In a young haustorium the contents are invariably dense with a high population 
of ribosomes, a profuse system of endoplasmic reticulum and relatively few vacuoles 
(Chou, 1970). The dictyosomes occur more frequently and the mitochondria are 
strikingly irregular (Figs. 11, 12, 15B). The same pattern of these structures are also 
present in young penetration hyphae. A complicated membrane system of unknown 
nature and origin is always present (Fig. 16B, C). One type consists of a complicated 
system of tubules and vesicles enclosed by a unit membrane. The inter- tubular spaces 
do not contain ribosomes. This organelle looks like a lomasome except that there is no 
apparent connection with plasmalemma. Another type consists of whorls of closely 
packed membranes formed in vacuoles (Fig. 16C). Generally the lomasomes are more 
or less hemispherical to saucer-shaped, about 0.2-0.3// in the longer diameter, but 
occasionally they can extend to 2-3// in diameter (Fig. 12). The tubules and vesicles of 
lomasomes range from 15 to 80//m in diameter. The nuclei are about 3-3.5// in 
diameter. As many as three sections of nuclei are observed in one haustorium section 
(Fig. 15A). The nuclei envelope consists of a double membrane interrupted by pores. 
The envelope is very similar in form to the endoplasmic reticulum and connections 
between these two are often observed. The endoplasmic reticulum is mainly of the 
smooth type (Fig. 15A, 12) enlarged in part to form cisternae of various forms. The 
mitochondria are large (1-2// in diameter), usually elongated dumb bell shaped or 
irregularly branched (Figs. 12, 15A, 7A, 9D). Those in old haustoria are roundish with 
a much less dense matrix (Fig. 15B). 

iii) The host - pathogen interface: The external surface of the haustorial walls 
always appears to consist of very electron dense layer (Figs. 10B, 11, 12, 15A, 16A, 17C, 
D) which is well-developed at the earliest stage of haustoria development (Chou, 1970). 
The outer region of the hyphal wall can be further differentiated into a well-defined, 
very dense and thin outer boundary, about 50-100A thick, and an inner less dense zone 
of rather obscure lateral limit (Fig. 14E). The hyphal wall thus appears to be a three- 
layered structure. The zone of apposition of the haustorial wall consists of a well- 
defined very dense and thin outer layer approximately 50- 100 A thick and a broad, 
slightly less-dense inner zone without a well-defined boundary (Fig. 14D). Chou (1970) 
proposed that the zone of apposition should be termed as an outer and inner dense zone 
being both an integral part of the haustorial wall. The surface of the haustorium neck 
is covered by a dense layer much thicker than that of the rest of the haustorium. Its 
surface always appears to be deeply roughened with numerous vesicular and tubular 
extensions (Figs. 12, 13B, 17B). Dense granular bodies can sometimes be seen lodged 
between the surface of the dense layer and the invaginated host plasmalemma and in 
both of the matrix of the dense layer and of the tubular extension (Figs. 13A, 17B). 
There is a frequent occurrence of a porous substance of uniform pore diameter (about 



32 




Fig. 1 1 . Electron micrograph of a section of a haustorium in host mesophyll cell at 6h after inoculation, x 1 2000 (Reprinted from 
C.K. Chou. 1970. An electron microscope study of host penetration and early stages of haustorium formation of 
Peronospora parasitica (Fr.) Tul. on cabbage cotyledons. Ann. Bot. 34: 189-204, by permission of the author and the 
publisher Academic Press Limited, London). 



33 




Fig. 1 2. Electron micrograph of a section of a haustorium in host mesophyll cell at 6h after inoculation, showing the sac-like sheath 
and numerous vesicles (arrow pointed) and intravacuolar vesicles (pointed out by double arrow) in the sheath matrix, x 
8580 (Reprinted from C.K. Chou. 1970. An electron microscope study of host penetration and early stages of haustorium 
formation of Peronospora parasitica (Fr.) Tul. on cabbage cotyledons. Ann. Bot. 34: 189-204, by permission of the author 
and the publisher Academic Press Limited, London). 



34 



ryg"*;- sin 




Fig. 13. (A) Electron micrograph of a section of part of haustorium neck and sheath. X 24600; (B) Electron micrograph of a section 
of part of haustorium neck and sheath showing numerous vesicles (arrow pointed) and dense granules in the sheath matrix 
(Smx) and the dentate extensions (pointed out by double arrow) of the dense zone (z) of haustorium wall, x 33000; (C) 
Electron micrograph of a section of the interface between haustorium and host cytoplasm showing a dense vesicle (arrow 
pointed) like the secretory body, x 33000; (D) Electron micrograph of a section of haustorium sheath showing incorporation 
of host cytoplasm (arrow pointed) in the sheath matrix and numerous membrane-bounded vesicles both in host cytoplasm 
and the sheath matrix, x 33000 (Reprinted from C.K. Chou. 1970. An electron microscope study of host penetration and 
early stages of haustorium formation of Peronospora parasitica (Fr.) Tul. on cabbage cotyledons. Ann. Bot. 34: 189-204, 
by permission of the author and the publisher Academic Press Limited, London) 



35 




Fig. 14. (A) Electron micrograph 
of a section of haustorium 
in host mesophyll cell 
showing the vacuoles or 
provacuoles possibly in the 
process of fusion with each 
other and also with the 
sheath (arrow), x 7200; (B) 
Electron micrograph of a 
section of the interface 
between haustorium and 
host cytoplasm showing 
vesiculation of the host 
plasmalemma. x 48000; 
(C) Electron micrograph of 
a section of haustorium in 
host mesophyll cell 
showing fusion of 
vacuoles in host cytoplasm 
and sheath formation, x 
7200; (D) Electron 
micrograph of a section of 
interface between 

haustorium and host 
cytoplasm showing the 
structure of outer dense 
zone of haustorium wall 
distinguished into two 
well-defined layers (zl) 
and (z2). x 49500; (E) 
Electron micrograph of a 
section of intercellular 
hyphae showing the 
hyphal wall also exhibit- 
ing a dense outer layer composed of zl and z2. x 33000 (Reprinted from C.K. Chou. 1970. An electron microscope 
study of host penetration and early stages of haustorium formation of Peronospora parasitica (Fr.) Tul. on cabbage 
cotyledons. Ann. Bot. 34: 189-204, by permission of the author and the publisher Academic Press Limited, London). 



36 




Fig. 15. (A) Electron micro- 
graph of a section of 
haustorium in 

epidermal cell at 6h 
after inoculation 

showing the typical 
fine structure of 
haustorium at this 
stage. Ring formation 
in mitochondria 

pointed out by arrow, 
x 13200; (B) Electron 
micrograph section of 
haustorium in 

epidermal cell 45h 
after inoculation, x 
24000 (Reprinted 
from C.K. Chou. 1970. 

An electron microscope study of host penetration and early stages of haustorium formation of Peronospora parasitica 
(Fr.) Tul. on cabbage cotyledons. Ann. Bot. 34: 189-204, by permission of the author and the publisher Academic Press 
Limited, London). 



' 'A <- .»•••».* 




37 



Fig. 16. (A) Electron micrograph of a section of the interface between haustorium and host cytoplasm of mesophyll cell showing 
sphaerosome-like bodies (arrow pointed) in host cytoplasm, x 33000; (B & C) Electron micrograph of sections of young 
penetrating hyphae; (B) showing complicated membrane system of unknown nature. (C) showing intravacuolar membrane 
systems, x 55200 and 36000 respectively (Reprinted from C.K. Chou. 1970. An electron microscope study of host 
penetration and early stages of haustorium formation of Peronospora parasitica (Fr.) Tul. on cabbage cotyledons. Ann. 
Bot. 34: 189-204, by permission of the author and the publisher Academic Press Limited, London). 



38 

200A) covering the entire haustorium surface. The host plasmalemma covering the 
haustorium surface is often masked due to the accumulation of this substance (Fig. 17 A, 
D). 

As soon as the haustorium penetrates the host, the haustorium becomes covered 
with a layer, moulded to its shape and produced by the host protoplast (Fraymouth, 
1956). This layer is named " The Sheath " and appears to be composed of modified 
cellulose and callose. A sudden increase in the growth rate of the fungus often causes 
the sheath to burst, remaining as a collar around the base. A morphologically 
analogous structure enveloping a haustorium initial which has penetrated the host wall 
(Figs. 9C, 10A) has been detected in the cabbage - Peronospora system by Chou (1970). 
In mature haustoria which have differentiated into a neck and head, the sheath 
remains as a collar-like structure at the base (Figs. 11, 12) although completely 
ensheathed mature haustoria are sometimes observed under the light microscope (Fig. 
7D). Electron microscope observation revealed that the sheath is a sac-like structure 
sometimes flattened to a narrow strip (Fig. 11), but in most cases dilated to a broadly 
conical shape, and quite distinct in texture and electron density from the host wall. The 
sheath is bounded by a unit membrane which is generally presumed to be the host 
plasmalemma. No membranous structure can be detected along the sheath/host-wall 
interface, though the two can be clearly distinguished by their difference in electron 
density and texture. The sheath matrix is electron transparent, while the host wall is 
moderately electron dense and often exhibits a fibrillar structure (Fig. 8A). The sheath 
matrix is always permeated by large numbers of blurred electron-dense granules, and 
dense vesicles with single or double membranes. These vesicles appear to be of host 
origin, as they are also found in the adjacent host cytoplasm (Fig. 13B, D, 12). The 
sheath matrix is also interspersed with host cytoplasm (Fig. 13D) which occurs in 
isolated packets or as an extension of the adjoining host cytoplasm. The permeation of 
vesicles into the sheath matrix and the extension of host cytoplasm within it suggest 
a liquid or semi-liquid state of the sheath matrix (Fraymouth, 1956; Chou, 1970). 
During penetration, the host cytoplasm adjoining the sheath increase markedly in 
amount and comes to contain a large number of vacuoles (Figs. 11, 12, 14A, C) (Chou, 
1970). At an early stage of haustorium development, coalescence of these vacuoles with 
the sheath can be seen. Intrusion intravacuolar vesicles in the host cytoplasm and in 
the sheath can be seen (Figs. 12, 13A, C). 

iv) Conidiophore development: Conidiophore development of P. parasitica can 
be divided into five stages (Davison, 1968). 

(a) Conidiophore primordia: The emergence of P. parasitica from the host 
cotyledons during sporulation, can be seen as a densely stained region beneath 
the host stomata. In the substomatal space, a hyphal branch, about 5 /x in diam., 
grows towards the stoma and then between the guard cells (Fig. 18A). When the 



39 




Fig. 17. (A) Tangential section of 
the dense zone of 
haustorium neck showing 
foldings of host plasma- 
lemma (arrow pointed) 
forming tubular extens- 
ions and incorporation of 
numerous dense granules 
(d). x 33000; (B) Electron 
micrograph section of 
haustorium in host 
epidermal cell showing 
lomasome. x 55200; (C) 
Electron micrograph of a 
section of haustorium in 
host epidermal cell 
showing pinocytotic 

vesicles formed from host 
plasmalemma and 

abundant porous substance 
(arrow pointed) at the host- 
parasite interface, x 73200; 

(D) Electron micrograph section of interface between dense zone of haustorium and host cytoplasm showing the deposition 
of porous substance (arrow pointed), x 48000 (Reprinted from C.K. Chou. 1970. An electron microscope study of host 
penetration and early stages of haustorium formation of Peronospora parasitica (Fr.) Tul. on cabbage cotyledons. Ann. 
Bot. 34: 189-204, by permission of the author and the publisher Academic Press Limited, London) 



40 




] 



Fig. 18. (A) Section of wax- 
embedded material 
showing a hyphal branch 
growing towards a stoma; 
(B) section of wax- 
embedded material 
illustrating two conidio- 
phore primordia one of 
which is beginning to 
grow; (C) stained and 
macerated preparation of 
an unbranched conidio- 
phore; (D) stained and 
macerated preparation of 
a branched conidiophore; 

(E) a branched conidiophore with small spores in a stained and macerated preparation; (F) very young spores; (G) mature 
spores; (H) mature spores delimited by a cross wall (arrow); (1-L) frames from the cine film illustrating the development 
of conidiophores A, B, and C; (I) incubation time 3h 30 min.; (J) incubation time 3 h 50 min.; (K) incubation time 4h 10 
min.; and (L) incubation time 4h 30 min. A-H scale line is 10m, I-L scale line is 100m (Reprinted from E.M. Davison. 
1968. Development of sporangiophores of Peronospora parasitica (Pers. ex Fr.) Fr. Ann. Bot. 32: 623-631, by permission 
of the author and the publisher Academic Press Limited, London). 




41 

tip of this hypha is about level with the top of the guard cells it becomes more rounded, 
and completely blocks the stomatal pore (Fig. 18B). It is referred to as conidiophore 
primordium. According to Shiraishi et al. (1975) a contracted image is found in the 
region when the conidiophores develop. 

(b) Unbranched conidiophores: It is the earliest stage of conidiophore 
development visible on the surface of the host, and can be seen about 4h after the 
cotyledons are placed in a moist, dark environment. From the primordia, 
unbranched conidiophores may develop immediately or a narrow wall surrounding 
a "blow out" forms at the apex (Fig. 18B). The basal constriction surmounted by 
a bulge which is observed in older conidiophores are probably the result of the 
"blow-out" formation. Developing conidiophores are more or less cylindrical at this 
stage (Fig. 18C), approximately 10-12 fx in diameter and of varying length. 

(c) Production of branches: When the conidiophores reach about two-third 
of its eventual height, branches are formed one at a time, just behind the 
conidiophores apex (Fig. 18D). Secondary and tertiary branches are also formed 
which are narrower than the primary ones. The conidiophore axis is also narrower 
at the apex, with decrease in branch diameter being proportional to the increase 
in branch length. The ultimate branches are very slender, often about 1 fx in 
diameter, and usually curved. Branches form at a projected angle of 55-85° with 
the major axis, and the number produced is approximately proportional to the 
condiophore height. 

(d) Development of conidia: Young conidia are formed about 2h after initiation 
of branch production. Initially, conidia are spherical, but as they increase in size 
they become ellipsoidal (Fig. 18E). Conidia produced on a single conidiophore are 
of uniform size, but conidia borne by different conidiophores frequently vary in 
size. 

(e) Formation of a cross wall: Conidia are delimited by a cross wall about 2h 
after the beginning of conidial formation, when they reach about 15 x 20 fu. in size 
(Fig. 18H). However, the cross walls are observed only occasionally since spores 
are usually detached before cross wall formation. 

v) Conidiophore growth: The increase in conidiophore length shows an initial 
slow period of elongation, just after the fungus has emerged from the host leaf followed 
by a rapid increase (Figs. 19, 20, 21). During branch formation increase in length is 
slightly slower and less regular, while just before spore formation conidophore 
elongation slows down and almost ceases. Once formed, the spores enlarge rapidly, but 
increase in conidiophore length is only by the enlargement of the apical spore. The most 
rapid rate of elongation of conidiophores is 100-200 fJh. As the branches usually begin 



42 



::- ■ 






Vr" 




/ 







-v-— * — — 




Fig. 19. Continued development of conidiophores (A) incubation time 4h 50 min.; (B) incubation time 5h 10 min.; (C) incubation 
time 5h 30 min.; (D) incubation time 5h 50 min.; (E) incubation time 6h 10 min.; (F) incubation time 6h 30 min.; (G) 
incubation time 6h 50 min.; and (H) incubation time 7h 30 min. Scale line is 100^ (Reprinted from E.M. Davison. 1968. 
Development of sporangiophores of Peronospora parasitica (Pers. ex Fr.) Fr. Ann. Bot. 32: 623-631, by permission of the 
author and the publisher Academic Press Limited, London). 



43 



500 



400 



300 - 



a. 
c 






£ 200 - 



100 - 



...J- . 



*"• 



• > 




_L 



_l_ 



_L 



4 5 6 7 

Incubation time in hours 



Fig. 20. Increase in length of five individual conidiophores growing in the humidity chamber, br: time at which branching 
commenced; sp: spore formation (Reprinted from E.M. Davison. 1968. Development of sporangiophores of Peronospora 
parasitica (Pers. ex Fr.) Fr. Ann. Bot. 32: 623-63 1 , by permission of the author and the publisher Academic Press Limited, 
London). 



44 




35 40 45 50 55 60 65 
Incubation time in hours 



70 75 



80 



Fig. 21. Increase in length of conidiophore s A, B and C. or: formation of primary branch; sp: spore formation (Reprinted from EM. 
Davison. 1968. Development of sporangiophores of Peronospora parasitica (Pers. ex Fr.) Fr. Ann. Bot. 32: 623-631. by 
permission of the author and the publisher Academic Press Limited, London). 



45 

about two third of the way up the final length of the conidiophore stalk, late 
conidiophores are usually shorter, less profusely branched, and bear fewer spores than 
the conidiophores formed earlier. Although increase in volume may be approximately 
linear during the growth of unbranched conidiophores and branch production, there is 
a decrease in the rate of volume increase just before spore formation. This is followed 
by a massive increase in volume just after spore formation, when the total 
conidiophores volume may be more than quadrupled (Fig. 22) depending on the number 
of spores produced. The inflation of the branch apex is a gradual process which occurs 
without any interruption (Fig. 23) in branch elongation (Davison, 1968b). 

vi) Conidial formation: Surface ultrastructure of conidia, germ tubes, 
appressoria and conidiophores of P. parasitica infecting Japanese radish has been 
observed by Shiraishi et al. (1974) through scanning electron microscopy (Figs. 24-27). 
Mature conidia are approximately 7 yum in width and 10 nm in length. Conidia are 
formed directly from the swelling tips of the conidiophores, and they have the same 
surface structure as the conidiophores. Old conidia have many wart-like structures, 
although the mature conidiophores have a smooth surface (Figs. 26, 27). 

vii) Host response: The host protoplast of epidermal and mesophyll cells respond 
differently to infection by the downy mildew pathogen (Chou, 1970). The epidermal 
cells in most cases respond vigorously to infection resulting in a severe disruption of the 
protoplast. The central vacuoles contract and undergo fragmentation. The 
plasmalemma is broken down or detached from the wall and numerous vesicles are 
formed from it. The cytoplasm, which originally appeared as a thin peripheral coating 
of the wall, is either dislocated and aggregated into a vacuolated blob or completely 
dispersed to the extent that its identity cannot be discerned. Apparently intact 
mitochondria and chloroplasts appear to be set free from the groundplasm. 
Consequently, haustoria in epidermal cells are not surrounded by a clearly defined layer 
of host cytoplasm. Haustoria formation in a mesophyll cell is less disruptive. The host 
cytoplasm is invaginated by the invading haustorium while the tonoplast and 
plasmalemma remain intact. 

viii) Cytology and genetics: The haploid chromosome number of P. parasitica 
is n = 18-20 and it is a tetraploid (Sansome and Sansome, 1974). Nuclei, mitochondria, 
lipid material, protein and RNA in the intercellular mycelia and haustoria of P. 
parasitica are uniformly distributed (Davison, 1968a). Insoluble carbohydrate material 
has been detected in the fungal cell wall. Callose sheaths are occasionally seen 
partially surrounding the haustoria. A distinct plasmalemma, porate nuclei, tubular 
endoplasmic reticulum, mitochondria with tubular cristae, golgi dictyosomes and lipid 
bodies are present within the protoplast (Ehrlich and Ehrlich, 1966). The distribution 
of organelles, storage products and other substances within the developing 
conidiophores of P. parasitica is very different from the distribution observed within the 



46 



14 












A 


12 












B 


10 














O 8 
I 


- 








/ / c 




c 

• 6 
E 

3 

c 
> 4 






«P 








2 








s p y 






r> ' 






i 


$ p 


i | 





35 40 45 



50 5-5 60 6-5 
Incubation time in hours 



70 7-5 



Fig. 22. Increase in volume of conidiophores A, B, and C. sp: spore formation (Reprinted from E.M. Davison. 1968. Development 
of sporangiophores of Peronospora parasitica (Pers. ex Fr.) Fr. Ann. Bot. 32: 623-63 1 , by permission of the author and the 
publisher Academic Press Limited, London). 




10 



20 30 

Time in minutes 



Fig. 23. Increase in branch length and apical diameter during spore formation. I: branch length; b: apical diameter (Reprinted from 
E.M. Davison. 1968. Development of sporangiophores of Peronospora parasitica (Pers. ex Fr.) Fr. Ann. Bot. 32: 623-63 1 . 
by permission of the author and the publisher Academic Press Limited, London). 



47 




Fig. 24. (See legend page 50a) 



48 




Fig. 25. (See legend page 50a) 



49 




Fig. 26. (See legend page 50a) 



50 




Fig. 27. (See legend page 50a) 



50a 



Fig. 24 Electron micrograph of conidia, germ tubes and initial period of Peronospora parasitica invasion on Japanese radish 
leaves. (A) Mature conidium. The surface is rough, with wart-shaped structure; (B) Separation of a mature conidium 
from its conidiophore; (C) An appressorium above a stoma, and a penetration peg into the stomatal cavity; (D) 
Enlargement of C. Wrinkly structures on an appressorium in the initial period of formation; (E) An appressorium 
over a stoma 48-h after germination; (F) Enlargement of E. Slight degeneration of the epidermal cells where the 
appresorium is in contact with the stomatal guard cells; (G) Cuticular invasion. Germ tube growing from the side of a 
spore; (H) Enlargement of G. The appressorium is quite contracted (Reprinted from M. Shiraishi, K. Sakamoto, Y. 
Asada, T. Nagatani and H. Hidaka. 1975. A scanning electron microscopic observation on the surface of Japanese 
radish leaves infected by Peronospora parasitica (Ft.) Fr. Ann. Phytopath. Soc. Japan 41:24-32, by permission of the 
authors and the publisher Phytopathological Society of Japan.) 

Fig. 25 Electron micrograph of initial period of Peronospora parasitica invasion on Japanese radish leaves. (A) Invasion 
through a junction between a stomatal guard cell and an auxiliary cell; (B) Cuticular invasion of an auxiliary cell. The 
germ tube is quite extended, but invasion does not depend on a stoma being present; (C) Enlargement of B. The 
viscous substance used by the appressorium to adhere to the epidermal cell wall is not very visible; (D) Enlargement of 
C. The germ tube and appressorium are clearly contracted, and circular traces of where the penetration peg has entered 
can be seen in the epidermal cell wall; (E) Cuticular invasion with a long germ tube; (F) Cuticular invasion through a 
short germ tube. Although the conidium is adjacent to a stoma, germination has occurred from the conidium wall on 
the side away from the stoma, and cuticular invasion is taking place (Reprinted from M. Shiraishi, K. Sakamoto, Y. 
Asada, T. Nagatani and H. Hidaka. 1975. A scanning electron microscopic observation on the surface of Japanese 
radish leaves infected by Peronospora parasitica (Fr.) Fr. Ann. Phytopath. Soc. Japan 41:24-32, by permission of the 
authors and the publisher Phytopathological Society of Japan.) 

Fig. 26. Electron micrograph showing development of conidiophores and conidia of Peronospora parasitica on Japanese 
radish leaves (A) Conidiophores invariably grow out of stomata, sometimes two at a time; (B) A conidiophore 
branching during the initial stage of new growth; (C) Surface of a conidiophore during the initial stage of new growth, 
with a wavy structure; (D) An extended conidiophore with appearance of a crimp at the base; (E) Initial stage of 
conidium formation. The tips of the conidiophore swell, forming conidia. The conidiophores and conidia have similar 
surface structures; (F) Clusters of conidia that have matured and begun to take on a tuft-like shape (Reprinted from M. 
Shiraishi, K. Sakamoto, Y. Asada, T. Nagatani and H. Hidaka. 1975. A scanning electron microscopic observation on 
the surface of Japanese radish leaves infected by Peronospora parasitica (Fr.) Fr. Ann. Phytopath. Soc. Japan 41:24- 
32, by permission of the authors and the publisher Phytopathological Society of Japan.) 

Fig. 27. Electron micrographs showing conidiophores and conidia of Peronospora parasitica on Japanese radish leaves (A) 
Conidiophores without conidia. The area at the top right is a relatively young diseased area, and exfoliation of epideral 
cell wax and cuticular material can be seen; (B) Diseased area with advanced signs of disease. Wrinkles have appeared 
in the epidermis of the diseased area, and open stomata can be seen; (C) Diseased area with many developed 
conidiophores; (D) Diseased area with advanced symptoms of disease. A crimp in the base of the conidiophore is 
visible. Yeast-shaped fungi area also present; (E) Stoma in a healthy area. It is formed of two stomatal guard cells and 
several auxiliary cells; (F) The base of the conidiophore is crimped, perhaps due to mechanical force exerted by the 
stoma. Wrinkles on the surface of the host are clearly visible (Reprinted from M. Shiraishi, K. Sakamoto, Y. Asada, T. 
Nagatani and H. Hidaka. 1975. A scanning electron microscopic observation on the surface of Japanese radish leaves 
infected by Peronospora parasitica (Fr.) Fr. Ann. Phytopath. Soc. Japan 41:24-32, by permission of the authors and 
the publisher Phytopathological Society of Japan). 

Abbreviations for Figures 24-27 

A = Appressorium; C = Conidium; CP = conidiophore; DT = Diseased tissue; E = Epidermal cell wall; G = Germ tube; 
GC = Guard cell; HT = Healthy tissue; J = Junction line of the epidermal cell wall; S = Stoma. 



51 

intercellular mycelium (Davison, 1968c). In developing conidiophores, the nuclei, 
mitochondria, protein and lipid material are more or less uniformly distributed at first, 
but gradually shift into the conidia so that by maturity all these substances have 
relocated, leaving the conidiophores stalk and branches almost completely empty (Figs. 
28-30). Glycogen has not been detected within conidiophores or conidia of P. parasitica. 
Trehalose and either glucose or mannose are identified in the conidiophores and conidia 
of P. parasitica but sugar alcohols are absent. 

d. Physiologic specialization (Pathogenic variability) 

r 

Specificity in the downy mildew fungus on crucifers is very complex since it occurs 
on a wide range of wild hosts as well as agricultural and horticultural species. For most 
of these there has been little sustained effort to introduce resistance to the disease, and 
hence there has been less selection pressure exerted on the pathogen population than 
is the case with many other obligate parasites. Further impetus has been added by the 
exponential growth in research on the wild crucifer Arabidops is thaliana, as a host for 
P. parasitica, and serving as a model system for genetic and molecular analysis (Uknes 
et al., 1992). Discontinuities in the host range of isolates from different host genera and 
species suggest that the fungus may exist as a series of pathotypes adapted to each host 
of origin, although some cross-infection may occur. There is also growing evidence that 
within host species, specificity may be determined by genotype specific interactions 
consistent with a gene for gene recognition system (Lucas et al., 1988, 1994; Nashaat 
et al., 1995). Specificity might therefore be expressed at several levels including family, 
genus, species and cultivar or accession. In view of the close cytogenetic relationship 
between the major Brassica species, coupled with the strongly outbreeding nature of 
several of these, some overlap in the host range of species-adapted isolates is perhaps 
predictable. 

At the generic level, pathogenic specialization has been observed by several 
workers all around the world. Gardener (1920) and Kobel (1921) suggested that P. 
parasitica is highly specialized and seldom occurs in the same biological form on more 
than one crucifer. An isolate of P. parasitica obtained from turnip is able to infect 
seedlings of turnip but not rutabaga or radish (Gardner, 1920). In Holland disease on 
cabbages is classified in two groups, both representing distinct biological forms of the 
fungus. The first is characterised by short, ellipsoid conidia, and the second by larger, 
elongated conidia with protuberant apices. The average dimensions of the later group 
are 32.51 x 25.66 /u. and that of the former 26.67 x 23.13 ix, the corresponding ratios of 
length to breadth being 1.26 and 1.11, respectively (Thung, 1926). However, Gaumann 
(1926) sub-divided P. parasitica (= P. brassicae) into three biological strains, namely: 
1. f. sp. brassicae, the chief hosts of which are B. oleracea, B. napus, B. rapa, B. nigra, 
B. juncea, B. tournefortii and B. fructiculosa, but can also cause some infection on 
Sinapis arvensis, S. alba, Raphanus raphanistrum, R. sativus and Eruca sativa; 2. f. sp. 



52 



(a) Nuclei 



X?^ 





5? DQ%r x%c 




(b)RNA 






xj^c xy^ 



(c) Mitocliondru 





33%I XD%I XJ%? 



Fig. 28. The distribution of (a) nuclei; (b) RNA; and (c) mitochondria in the developing conidiophores of Peronospora parasitica 
(Reprinted from E.M. Davison. 1968. The distribution of substances in the sporangiophores of Peronospora parasitica (Pers. 
ex Fr.) Fr. Ann. Bot. 32: 633-647, by permission of the author and the publisher Academic Press Limited, London). 



53 







Fig. 29. Migration of nuclei (n) in to the anucleate spores of Peronospora parasitica (Reprinted from E.M. Davison. 1968. The 
distribution of substances in the sporangiophores of Peronospora parasitica (Pers. ex Fr.) Fr. Ann. Bot. 32: 633-647, by 
permission of the author and the publisher Academic Press Limited, London). 



54 



la) Lipid material 



%• 



^XP^x x%r 




(h) Protein 



OT^c 



^ 




X^XJ^c X%r JO^n 



M Invilublc 
i.irbohvd rates 




| 

kr XP^l 30%r X%r 



Fig. 30. The distribution of (a) lipid material; (b) protein; and (c) insoluble carbohydrates in the developing conidiophores of 
Peronospora parasitica (Reprinted from E.M. Davison. 1968. The distribution of substances in the sporangiophores of 
Peronospora parasitica (Pers. ex Fr.) Fr. Ann. Bot. 32: 633-647, by permission of the author and the publisher Academic 
Press Limited, London). 



55 

sinapidis, the principal hosts of which are S. arvensis and S. alba, but is also able to 
produce sub-infections on all the above mentioned species of Brassica (except B. rapa 
and B. juncea) and Raphanus, with occasional conidiophore formation on B. oleracea; 
and 3. f. sp. raphani, the chief hosts of which are R. raphanistrum and P. sativus, but 
can also produce sub-infections on all the above mentioned species of Brassica (except 
B. fructiculosa) , as well as on S. arvensis and S. alba, with occasional conidiophore 
formation on B. oleracea and B. napus. The downy mildew on radish does not attack 
cabbage (B. oleracea var., bullata and capitata) and is slightly pathogenic on Chinese 
cabbage (B. pekinensis, B. chinensis), rape (B. campestris) and mustard (B. juncea) 
(Hiura and Kanegae, 1934). Conversely, the form derived from B. pekinensis does not 
infect radish but is allied to one on B. chinensis and rape. The forms derived from B. 
pekinensis, B. chinensis and rape are mutually pathogenic on one another (Hiura and 
Kanegae, 1934). 

Wang (1944) classified the reaction of the hosts into four categories: 1. susceptible 
with normal symptoms; 2. resistant, showing large necrotic spots; 3. para-immune, 
showing slightly visible necrotic dots; and 4. immune, with no visible symptoms. Three 
pathotypes of P. parasitica were differentiated: P. parasitica Brassicae on Brassica, P. 
parasitica Raphani on Raphanus and P. parasitica Capsellae on Capsella. The three 
pathotypes were not mutually compatible with each other's host. Six forms of P. 
parasitica Brassicae were differentiated by their reaction to B. chinensis, B. oleracea, 
B. juncea and B. napobrassicae. Wang (1944) prepared a dichotomous key to 
physiological forms from China as follows: 

A. Capsella bursa-pastoris, immune 

B. Raphanus sativus, immune or para-immune variety Brassicae 

C. B. oleracea, resistant or para-immune 

D. B. chinensis, susceptible 

E. B. juncea (Meitan Dav Yu Tsai), susceptible Ph.fm. 1. 
EE. B. juncea (Meitan Dav Yu Tsai,) resistant 

F. B. napobrassica, immune Ph. fm. 2. 
FF. B. napobrassica, resistant Ph. fm. 3. 
DD. B. chinensis, resistant 

E. B. juncea (Dav Ching Tsai), susceptible Ph. fm. 4 

EE. B. juncea (Dav Ching Tsai), resistant Ph. fm. 5 

CC. B. oleracea, susceptible Ph. fm. 6 

BB. R. sativus, susceptible variety Raphani 

AA. C. bursa-pastoris, susceptible variety capsellae 

Felton and Walker (1946) and Natti (1958) differentiated the races of P. parasitica 
found on R. sativus, and B. oleracea on the basis of their host specificity. Morris and 
Knox-Davies (1980) also indicated distinct races of P. parasitica on B. oleracea and R. 



56 

raphanistrum based on host specificity. 

P. parasitica f. brassicae on cabbage, f. rapae on turnip, f. rapiferae on B. rapa, f. 
rapifera, f. napi on rape, f. raphani on radish, and f. sinapidis on sinapis alba have been 
distinguished as special forms of P. parasitica from Leningrad though all are similar 
morphologically (Dzhanuzakov, 1963). 

Three vars. of P. parasitica have been differentiated in 35 samples of downy 
mildew from B. pekinensis and other crucifers, namely f. sp. brassicae on Brassica, f. sp. 
raphani on Raphanus, and f. sp. capsellae on Capsella (Chang et al. 1964). P. parasitica 
f. sp. Brassicae exists in at least 3 different subforms (pekinensis, oleracea and juncea). 
Isolates from B. pekinensis, B. chinensis and turnip were classified in the same group 
and can attack all three hosts but did not infect Capsella bursa-pastoris, radish, 
cabbage, Chinese mustard, and B. juncea var. multiceps. B. juncea var. megarrhiza 
expressed various reactions to these isolates. Isolates from Chinese mustard, B. juncea 
var. megarrhiza, and B. juncea var. multiceps were limited to these hosts, except that 
those from Chinese mustard did not infect some vars. of B. pekinensis, B. chinensis and 
turnip. Radish isolates are of two types, one infects only radish, the other vars. of B. 
pekinensis, cabbage and turnips as well as radish. Isolates from cabbage and C. bursa- 
pastoris are host specific. 

In Norway, cross-inoculation experiments with downy mildew from cabbage, turnip 
rape and radish indicated the occurrence of different races on cabbage and radish 
(Semb, 1969). 

According to Natti et al. (1967) the predominant physiologic race of P. parasitica 
pathogenic to broccoli and other types of B. oleracea grown commercially in New York 
were race 1 and race 2. The later race was pathogenic to plants resistant to race 1. 

Dickinson and Greenhalgh (1977) observed a wide variation in the reaction of 
seedlings of different crucifers species to isolates of Peronospora derived from Brassica 
and Raphanus species (Table 2). 

In India, P. parasitica isolates from different hosts vary in host range. Isolates 
from Brassica, Raphanus, Eruca and Sisymbrium are not cross infective (Bains and 
Jhooty, 1983). Recently Mehta and Saharan (1994) tested the host range of 9 isolates 
of P. parasitica collected from the leaves and stagheads of 6 host species on 17 host 
differentials (Tables 7, 8). Isolates from brassica oilseeds infected all species, except B. 
alba, whereas isolates from cauliflower leaves do not infect B. carinata, B. alba, B. 
nigra, B. chinensis, B. pekinensis and B. napus (Table 8). There was no significant 
differences among the conidial size of the isolates collected from leaves and stagheads, 
but significant differences were observed among these groups (Table 9). The isolates 



57 



Table 7. List of host differentials (Mehta and Saharan, 1994) 



Common name 


Species 


Cultivar 


Indian Mustard (Raya) 


Brassica juncea 


RH-30 


Toria 


Brassica campestris var. toria 


TH-68 


Yellow sarson 


Brassica campestris var. yellow sarson 


YSPB-24 


Brown sarson 


Brassica campestris var. brown sarson 


BSH-1 


Ethiopian mustard 


Brassica carinata 


HC-1 


White mustard 


Brassica alba 


Local 


Black mustard 


Brassica nigra 


Local 


Chinese mustard 


Brassica chinensis 


Local 


Chinese mustard 


Brassica pekinensis 


Local 


Rapeseed 


Brassica nap us 


GSL-1 


Wild turnip 


Brassica tournefortii 


Local 


Cabbage 


Brassica olercea var. capitata 


Pride of India 


Cauliflower 


Brassica oleracea var. botrytis 


Snowball- 16 


Turnip 


Brassica rapa 


White Purple Top 


Knol Khol 


Brassica caulorapa 


Early White Vienna 


Taramira 


Eruca sativa 


Local 


Radish 


Raphanus sativus 


HR-1 



were classified into two distinct pathotypes, one from cauliflower and other from 
oilseeds brassica. There was no significant difference between the isolates in 
percentages of spore germination (Table 10). 



Specific populations of P. parasitica differing in pathogenesis and host specificity 
were reported from Bulgaria (Masheva et al., 1996a, b). The populations formed at a 
lower temperature were more aggressive on cabbage heads. 

In the United Kingdom, differential host resistance in relation to pathogenic 
variation of isolates derived from the same host species were identified in B. rapa (Moss 
et al., 1991; Silue et al., 1996), B. napus (Nashaat and Rawlinson, 1994), B. juncea 
(Nashaat and Awasthi, 1995) andB. oleracea (Silue et al., 1996). Isolates from different 
Brassica species found to be most virulent on their species of origin, were nevertheless 
able to grow to less extent on other Brassica species (Sheriff and Lucas, 1990). Nashaat 
and Awasthi (1995) identified five groups of B. juncea accessions with differential 
resistance to U.K. isolates Rl and P003 derived from oilseed rape (B. napus ssp. 
oleifera) and Indian isolates IP01 and IP02 derived from mustard {B. juncea) (Table 11). 
All B. juncea accessions were resistant to isolates from B. napus, but at the same time 
B. napus cv. Arian was resistant to isolates from B. juncea. Twenty-one differential 
responses to P. parasitica isolates from B. oleracea and two from B. rapa were 
identified. Of the seven isolates tested, four were from crops of cauliflower in France, 



58 



Table 8. Response of seventeen Brassica species to nine isolates of 
Peronospora parasitica (Mehta and Saharan, 1994) 



Toria Yellow 

Differential 

Hosts (TO YSj 



Sources of P. parasitica isolates/reaction 
Sarson Brown Sarson Raya Cauliflower B. nigra 



YS, 



BSj 



BS, 



R> 



K , 



c x 



(BN 2 ) 



B. campestris 

var. toria +(4 
B. campestris 

var. Y. Sarson +(6 
B. campestris 

var. B. Sarson +(4 

B. juncea +(4 

B. carinata +(6 
B. alba 

B. nigra +(6 

B. chinensis +(6 

B. pekinensis +(6 

B. napus +(4 

B. tournefortii +(6 

Eruca sativa +(4 
Raphanus 

sativus +(4 
B. oleracea 

var. capitata +(4 
B. oleracea 

var. botrytis +(4 

B. rapa +(4 

B. caulorapa +(4 



•) +(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


) +(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


) +(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


) +(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


) +(6) 


+(4) 


+(6) 


+(4) 


+(4) 


+(4) 


) +(6) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


) +(6) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


) +(6) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


) +(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


) +(6) 


+(4) 


+(6) 


+(4) 


+(4) 


+(4) 


) +(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


) +(4) 


+(6) 


+(4) 


+(4) 


+(4) 


+(4) 


) +(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


) +(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


) +(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


) +(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 



+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


- 


+(4) 


. 


+(4) 


- 


+(4) 


- 


+(4) 


- 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 


+(4) 



( ) = Incubation period in days; + = Infection; - = No infection 
B = Brassica; 1 = Leaf inoculum; 2 = Inflorescence inoculum 



two from oilseed rape in the U.K. and one was from mustard in India. All Raphanus 
sativus accessions were resistant to all seven isolates (Silue et al., 1996). 



Differential resistance of rapid cycling and commercial genotypes of B. rapa were 
also reported from the U.K. (Moss et al., 1988). A range of differential host responses 
were characterized by four homologous isolates (Table 12). Pathogen isolates were also 
characterized in relation to their sexual compatibility type and response to phenylamide 
fungicides (Moss et al., 1988). 



59 



Table 9. Conidial size of Peronospore parasitica isolates derived from 
elevenBrassica species (Mehta and Saharan, 1994) 









Conidial dimensions (u) 




Source of 


P. parasitica 
Isolates* 


Range 


Average 


Isolates 


Length 


Width 


Length 


Width 


B. campestris 




19.50-29.25 


14.62-27.30 


25.93 


19.30 


var. toria 


r T l 










B. campestris 




21.45-29.25 


19.50-24.37 


25.35 


21.74 


var. Y. Sarson 


YS, 










B. campestris 




19.50-29.25 


19.50-24.37 


26.81 


23.39 


var. Y. Sarson 


YS 2 










B. campestris 




21.45-29.25 


19.00-24.37 


25.35 


21.45 


var. B. Sarson 


BSi 










B. campestris 




21.93-29.25 


19.50-26.81 


26.56 


23.64 


var. B. Sarson 


BS 2 










B. juncea 


Ri 


20.47-29.25 


19.50-26.32 


25.64 


21.84 


B. juncea 


R* 


19.50-29.25 


19.50-26.81 


27.05 


22.90 


B. oleracea 




19.50-29.25 


17.55-24.37 


23.30 


20.76 


var. botrytis 


Cx 










B. tournefortii 


BT 2 


19.50-24.37 


16.57-19.50 


23.59 


20.96 


Raphanus sativus 


RS 2 


19.50-24.37 


18.52-21.45 


22.03 


19.69 


B. nigra. 


BN 2 


21.93-29.25 


19.50-26.81 


26.81 


22.90 


CD 0.05 




- 


- 


2.43 


1.78 



* Source of inoculum: 1. leaves; 2. hypertrophied inflorescences 



Eleven isolates of P. parasitica tested on rapid cycling populations of B. rapa (aa, 
CrGc-1-1), B. nigra (bb, CrGc-2-1), B. oleracea (cc, CrGc-3-1), B. juncea (aabb CrGc-4-1), 
B. napus (aacc CrGc-5-1), B. carinata (bbccCrGc-6-1) and R. sativus (rr CrGc-7-1) 
indicated specificity towards particular genotypes within each rapid cycling population 
(Hill et al., 1988). 



Variation in the response of different host lines to P. parasitica has also been 
detected within wild crucifer species such as Shepherd's Purse (Capsella bursa-pastoris) 
and Arabidopsis thaliana (Lucas et al., 1994). 

Moss et al. (1994) attempted to cross fungal isolates originating from different 
Brassica spp. by co-inoculating host lines previously identified as susceptible to these 



60 



Table 10. Percent conidial germination of Peronospora parasitica isolates 
at 18 °C (Mehta and Saharan, 1994) 



Incubation 


cauliflower 


Toria 


Brown Sarson 


Yellow Sarson Raya 


period (h) 


«y 


(T,) 


CBS,) 


(YS,) (R,) 



0.5 














1.0 











10.57 


1.5 


12.20 


13.73 


15.72 


11.87 32.46 


2.0 


36.50 


33.25 


45.87 


39.31 52.68 


3.0 


66.30 


63.25 


69.12 


61.06 69.12 


4.0 


84.25 


83.86 


80.99 


81.92 88.85 


5.0 


86.02 


84.91 


82.61 


84.16 88.85 


6.0 


86.51 


87.36 


82.90 


85.42 89.19 




(68.72) 


(70.11) 


(65.80) 


(67.85) (73.20) 



LSD 0.05 = NS 



isolates. A proportion of oospore progeny recovered from these crosses appeared to be 
hybrids and had reduced virulence on both hosts of origin. The differential resistance 
to P. parasitica identified in Brassica spp. can be used for future studies of the genetics 
of the host-pathogen interaction and for breeding for disease resistance. 

Tham et al. (1994) used RAPD analysis to compare 16 isolates of P. parasitica from 
two different Brassica species, B. napus and B. oleracea. Two out of twenty random 
primers screened gave reproducible band patterns capable of discriminating between 
the different host-adopted isolates (Fig.31 ). 

e. Heterothallism and homothallism: Induction of the sexual process in several 
downy mildew species requires the presence of two strains of opposite mating type. 
Such heterothallic behaviour has been reported for P. parasitica (DeBruyn, 1937; 
McMeekin, 1960; Kluczewski and Lucas, 1983; Sherriff and Lucas, 1989b; Sequeira and 
Monteiro, 1996). Homothallic forms of P. parasitica have also been observed (De Bruyn, 
1937; Sherriff and Lucas, 1989b; Sequeira and Monteiro, 1996). The two forms of 
sexual reproduction are very important for the maintenance and evolution of the fungal 
strains and epidemiology of the disease. On B. oleracea and B. campestris heterothallic 
isolates of two mating types designated as P, and P 2 have been identified. Isolates from 



61 



LANES 



13 14-17 



kb 

3.0 

2.0 
1.6 

1.0 



0.5 




Fig. 31. Random amplified polymorphic DNA (RAPD) from 16 isolates of crucifer downy mildew (Peronospora parasitica). 
Lanes 2-13 are isolates from oilseed rape Brassica napus and lanes 14-17 are isolates from cauliflower B. oleracea 
(Reprinted from F.Y. Tham, J. A. Lucas and Z.A. Wilson. 1994. DNA fingerprinting of Peronospora parasitica, a 
biotrophic fungal pathogen of crucifers. Theoretical and Applied Genetics 88:490-496, by permission of the authors 
and the publisher Spnnger-Verlag New York Inc., U.S.A.). 



62 

Table 11. Sources of seed for accessions of Brassicajuncea, arranged in five 
groups according to the response of their seedlings at the 
cotyledon stage to Peronospora parastica and one accession of B. 
napus (Reprinted from N.I. Nashaat and R.P. Awasthi. 1995. 
Evidence for differential resistance to Peronospora parasitica 
(downy mildew) in accessions of Brassicajuncea (mustard) at the 
cotyledon stage. J. Phytopathology 143:157-159, by permission of 
the authors and the publisher Blackwell Wissenschafts-Verlag 
GmbH, Germany) 





Seed 




Seed 


B. juncea* 


source b 


B. juncea 


source b 


GROUP A 




Group C 




RES-BJ01 (Kranti) 


(India) 


Ecotype/BGRC 34263 


FAL 


RES-BJ02 (Krishna) 


(India) 


BGRC 34283 


FAL 


RES-BJ03 (Varuna) 


(India) 


BGRC 34291 


FAL 


RES-BJ04 (BGRC 34253) 


(FAL) 


BGRC 34781 


FAL 






BGRC 34789 


FAL 






BGRC 46069 


FAL 


GROUP B 








Chang Yang Huang Jie 


HAU 


BGRC 46071 


FAL 


BGRC 34294 


FAL 


PPBJ-1 
Skorosjelka-2/ 


India 


GROUP C 




BGRC 34275 


FAL 


Aurea/BGRC 28602 


FAL 


Stephniacka/ 




Blaze/BGRC 30288 


FAL 


BGRC 34274 


FAL 


Burgonde/BGRC 30289 


FAL 


Stoke/BGRC 51764 


FAL 


Commercial Brown 


AgCda 


Yi Men Feng Wei Zi 


HAU 


Cutlass 


AgCda 


Zaria/BGRC 16254 


FAL 


Ecotype BGRC 34255 


FAL 






GROUP D 




GROUP E 




Hatano/BGRC 22527 


FAL 


BGRC 34282 


FAL 


BGRC 34239 


FAL 






Landrace/BGRC 46323 


FAL 


B. napus 




Larja/BGRC 34273 


FAL 


Ariana 


Semundo 


Line/BGRC 34295 


FAL 







RES-BJ01 to RES-BJ05, lines selected from seedling population of accessions in 
parethesis. 



63 



Agriculture and Agri-Food Canada, Saskatoon Research Centre, Canada. 
FAL, Institut fur Pflanzenbau und Pflanzenziichtung, Braunschweig, Germany. 
HAU, Huazhong Agricultural University, Wuhan, P.R. China. Semundo, 
Semundo Ltd., Cambridge, UK. 



oilseed rape B. napus have been found to be uniformly homothallic and remained self- 
fertile even after months of laboratory subculture (Sherriff and Lucas, 1989b). In a 
cytogenetic study of heterothallic and homothallic isolates of P. parasitica at metaphase 
1 of meiosis, a ring of four chromosomes is found (Sherriff and Lucas, 1989a). This ring 
is interpreted as a reciprocal translocation complex between chromosomes carrying the 
mating type alleles. In homothallic isolates a fifth chromosome is associated with the 
ring of four. The self fertility of these isolates may therefore be due to the presence of 
a third mating type allele on the fifth chromosome, a condition known as secondary 
homothallism. The determination of sexual compatibility type (SCT) of an unknown 
isolate can be achieved by mixing conidia in a 1:1 ratio with isolates of known SCT and 
inoculating to a common compatible host. In heterothallic isolates, oospores may form 
in combination with isolates of opposite SCT. Isoenzyme markers are particularly 
important in discriminating between self and true hybrid progeny (Moss et al., 1988). 

f. Perpetuation 

Oospores formed in malformed and senesced host tissues constitute an important 
means of survival of P. parasitica over periods of unfavourable conditions (Gauman, 
1926; Kolte, 1985). It is also known to survive through mycelium and conidia (Jang and 
Safeeula, 1990b; Krober, 1970; McMeekin, 1969; Vishunavat and Kolte, 1993). 

i) Mycelium: The presence of P. parasitica mycelium in the seed coat of Chinese 
cabbage has been recorded by Chang et al. (1963). According to Jang and Safeeulla 
(1990c) the presence of mycelium in the pericarp and embryo of radish seeds varies from 
0.1 to 12.5% (Table 13, 14). The coenocytic branched mycelium is clearly visible in the 
intercellular space of the pericarp. In the embryonal tissues, the mycelium is 
comparatively thin. The percentage of seeds with viable mycelium is directly correlated 
with the percentage of embryo infection. 

ii) Conidia: Conidia of P. parasitica on cabbage survive longer under cool, dry 
conditions (Krober, 1981). Relative humidity is more important than temperature. In 
the field conidia can survive on detached leaves of Kohlrabi for 10 days during warm 
days. When buried in dry soil conidia can survive for 110 days. The survival period is 
greatly reduced to a maximum of 22 days if the soil is moist. Marked reduction in 
survival has also been observed after storage in both dry and moist soils during the 



64 

Table 12. Differential virulence of P. parasitica isolates from B. campestris 
on six hosts lines (Reprinted from NA Moss, I.R. Crute, J.A. Lucas 
and P.L. Gordon. 1988. Requirements for analysis of host-species 
specificity in Peronospora parasitica (downy mildew). Cruciferae 
NewsLetter 13:114-116, by permission of the authors and the 
publisher) 



Host Lines Reaction of P. parasitica isolates 

P007 P008 P013 P014 



+ + 

+ + 

+ + 
+ 



CA88014* 3 


+ 


JADE PAGODA 


+ 


CA87063* 


- 


SNOWBALL 


- 


CA87068* 


- 


CA87065* 


_ 



+ = susceptible, - = Resistant, a = universally susceptible, * = rapid cycling lines 



summer. The conidial viability is longest, up to 130 days, when the spores are stored 
in air-dried soil at a constant temperature of 5°C (Krober, 1970). Conidia kept at -25°C 
and relatively dry on leaf disks (air dried at 20°C) maintain a relatively high rate of 
germination after 1 year or longer. 

iii) Oospores: Ooospores formation is abundant in the infected tissues of all 
crucifers and they form primary source of survival for the pathogen (Le Beau, 1945; 
McMeekin, 1960; Chang et al., 1963; Kolte, 1985). In radish and rapeseed-mustard 
there is abundant production of oospores in infected leaf tissues, on the seed surface and 
pericarp and embryo of seeds (Jang and Safeeulla, 1990c; Vishunavat and Kolte, 1993). 
However, in rapeseed and mustard, seed transmission is low and may be nonsystemic, 
ranging from 0.4% to 0.9% in the seedlings grown from infected seeds (Vishunavat and 
Kolte, 1993). In radish seed transmission to the extent of 14% was observed by Jang 
and Safeeulla (1990d). 

iv) Axenic culture: P. parasitica hyphae grow on water agar from the infected 
tissues and form haustorium like structures (Ohguchi and Asada, 1989). The growth 
is greater on the modified Knop medium with many haustorium-like structures and 
conidiophores being formed on this medium. If cod-liver oil or minerals are added to 



65 

Table 13. Percentage seed infection by P. parasitica in R. sativus (Reprinted 
from P. Jang and K.M. Safeeulla. 1990c. Seed-borne nature of 
Peronospora parasitica in Raphanus sativus. Indian Acad. Sci. 
(Plant Sci.) 100:255-258, by permission of the authors and the 
publisher Indian Academy of Sciences) 





Place of collection 


Seed showing infection (%) 


Cultivar 


Pericarp 


Endosperm 


Embryo 


Japanese white 


Mysore Seed 
Multiplication Farm 


12.8 





12.5 


Arka nishant 


Indian Council of 
Agricultural Research 
Station, Bangalore 


0.5 





0.5 


Pusa desi 


Bangalore Seed Health 
Testing Station 


0.2 





0.3 


Pusa reshmi 


Bangalore Seed Health 
Testing Station 








0.1 



this medium then branched hyphae are formed. The fungus does not grow on Japanese 
radish root homogenate medium but grows well on the dialized homogenate medium. 
The decoction of residuum of the root homogenate and the sap in the intercellular 
spaces of the root tissues also stimulates the growth of the fungus. In the decoction 
medium the growth of the hypha is vigorous and the formation of conidiophores is 
stimulated. In the sap medium, the formation of a haustorium like structure is 
promoted (Ohguchi and Asada, 1989). 



g. Conidial discharge: The maximum conidial discharge of P. parasitica from 
Kohlrabi leaves is between 5-6 a.m. (258 conidia/cm 2 ) (Lin and Liang, 1974). The 
conidial discharge decreases greatly from 12 noon to 8 p.m. If infected leaves are 
covered with plastic during the night, then the production and discharge of conidia 
decreases drastically and the disease index is half that of uncovered seedlings. The 
discharge of conidia from diseased leaves of Chinese cabbage shows a periodic cycle each 
day (Fig. 32A) (Lin, 1981). Conidial release increases steadily after 2 a.m. each day, 
and reaches a peak around 6-8 a.m. Conidial discharge decreases rapidly after 8 a.m. 
Few conidia can be detected from noon to 10 p.m. The discharge of conidia is favoured 
by temperatures below 18°C and RH above 75% (Fig. 32B). If the conidia are ready to 



66 



300 




10 



i I I I I I — 1 V ! I I I I I I I I I I l_I L 



4 S 12 16 20 4 8 12 16 20 4 8 12 16 20 
IDoy IIDoy HI Day 

Time of day ( hr) 



55 



Fig. 32. (A) Pattern of Peronospora parasitica conidia discharge from infested Chinese cabbage plants; and (B) temperature and 
humidity on three fine days in November 1978 (Reprinted from C.Y. Lin. 1981. Studies on downy mildew of Chinese 
cabbage caused by Peronospora parasitica. Proc. First Intern. Symposium, Asian Vegetable Research Development 
Centre, Shanhua, Taiwan: pp. 105-1 12, by permission of the author and the publisher Asian Development Research 
Development Centre, Taiwan). 



67 

Table 14. Percentage of seedling infection by P. parasitica and seed 
transmission in R. sativus (Reprinted from P. Jang and K.M. 
Safeeulla. 1990c. Seed-borne nature of Peronospora parasitica in 
Raphanus sativus. Indian Acad. Sci. (Plant Sci.) 100:255-258, by 
permission of the authors and the publisher Indian Academy of 
Sciences) 



Seedling Seed Infection (%) 

Cultivar infection (%) Pericarp Embryo 



Japanese white 14.0 13.5 12.8 

Arka nishant 1.5 0.5 0.4 

Pusadesi 1.0 0.1 0.2 

Pusareshmi 1.0 0.0 0.1 



be released from (Fig. 33) conidiophores but the RH suddenly decreases, then the 
branched conidiophores become dry and the twirling movement of the drying 
conidiophores may flick the spores in to the air and discharge abundant conidia from 
the diseased leaf. According to Shao et al. (1990) conidia release during favourable 
temperature and RH conditions is three times higher in the morning than in the 
afternoon. 

Dispersal of conidia of P. parasitica on Lepidium virginicum begins with incipient 
desiccation and conclude with hygroscopic distortion of the aerial fructifications 
(Pinckard, 1942). Several complete twists occur in the portion of tall conidiophores 
extending up to the first branch, with a lesser number between each successively 
shorter branch. With the progress of drying, a twisting and binding motion is imparted 
to the sterigma-like structure on which the conidia are borne. If the process of 
desication stops, the twisting motion also stops. However, if humidity increases, the 
rotation reverses itself. Under conditions of delicate moisture balance, the breath of an 
observer is sufficient to induce the above mentioned movements. The outcome of the 
movement is the release of mature conidia. By slowly decreasing the vapour pressure, 
a point is reached when abscission occur, and the conidia are forcibly released with the 
stimulus for the requisite energy being derived from differential stresses set up within 
the sterigmata. The mechanical action of wind and rain, during periods of atmospheric 
saturation, does not appear to contribute significantly to dispersal of conidia. 



68 




( C) changes in comd.ophore on exposure to ^^^ZZ^onosporl parasitica. Proc. First Intern. 
Lin. 1981. Studies on downy m.ldew of Ch.nese cabbage caused oy / permission of the author 

Symposium, Asian Vegetable Research Development Centre, Shanhua, Ta.wan. pp. 
Ld the pub Usher Asian Development Research Development Centre, Ta.wan). 



69 

h. Conidial germination 

For germination of conidia collected from Chinese cabbage, 8-20°C is favourable 
with an optimum range of 12 - 16°C (Fig. 34) (Lin, 1981). Germ tubes usually grow 
normally and extensively at these temperatures. The germination rate of conidia is low 
and the germ tubes show limited and malformed growth at temperatures below 8°C and 
above 20°C. Conidia fail to germinate at extreme temperatures even after a long period 
of incubation. Conidial germination usually increases after treatment with hot water 
of up to 42°C (Fig. 35). Germination of conidia from Chinese cabbage (B. pekinensis) 
was optimum at 15 - 20 °C and was stimulated by light (Shao et al., 1990). 

P. parasitica sporulates on intact cabbage seedlings when incubated at 13°C or 
18°C in the presence of free water, or to atmospheric water potentials (i|/) of or -30 + 
10 bars (Table 15) (Hartman et al., 1983). The pathogen fails to sporulate at these 
temperatures when the atmospheric i|/ is -60, -90 or -120 bars. More conidia are 
produced at 13°C (1466 - 2265 conidia/45 mm 2 cotyledons) than at 18°C (821 - 1042 
conidia/45mm 2 cotyledon). Conidia germinate in the presence of free water but do not 
germinate when exposed to atmospheric i|r of 0, -30, -60, or -90 bars for 24 h. The level 
of atmospheric ij; and the presence or absence of free water during sporulation exerts 
preconditioning effects on the ability of conidia to germinate. 

Conidia collected from B. campestris (Toria, Brown sarson, Yellow sarson) and B. 
oleracea (cauliflower) leaves germinate after 1.5h at 18°C, whereas conidia derived from 
B. juncea germinate after lh (Mehta and Saharan, 1994). Germination increases as the 
incubation period is increased. For instance, more than 80% conidia germinate after 
4h (Table 10). 

i. Oospore germination 

The germination of oospores of P. parasitica infecting radish is dependent on 
temperature, light, pH of the medium, and age of oospores (Jang and Safeeulla, 1990a). 
The optimum temperature for germination is 23°C. Drying and chilling of oospores has 
no marked effect on germination. At a pH of 7.5 germination is 42% but at a pH of 4.5 
only 1% oospores germinate. Oospore germination also increases with age (Jang and 
Safeeulla, 1990a). 

4. INFECTION AND PATHOGENESIS 

Infection may be either general or local. In the former case, all or most of the 
leaves and inflorescence (which may be hypertrophied as a result of pre-infection with 
A. Candida) may bear conidiophores. Although some parts (especially the stem) may 
show no external injury, microscopic examination shows that the mycelia are in the 
tissues (Butler, 1918). Generalized infection is restricted to young tissues and this is 
why seedlings show completely infected leaves. 



70 



o 



!- 




-95 



-81 



-55 



73 

n 



< 



X 

c 

3 



-32 



Germination ( X ) 



Fig. 34. The effect of temperature and relative humidity on the germination of conidia of Peronospora parasitica (Reprinted from 
C.Y. Lin. 1981. Studies on downy mildew of Chinese cabbage caused by Peronospora parasitica. Proc. First Intern. 
Symposium, Asian Vegetable Research Development Centre, Shanhua, Taiwan: pp. 105-1 12, by permission of the author 
and the publisher Asian Development Research Development Centre, Taiwan). 



71 




36 30 3* 38 -42 *6 50 b* 58 62 

Water Temperature f*C 



Fig. 35. Effect of hot water treatment on the germination of (A) conidia of Peronospora parasitica, and (B) seeds of three Chinese 
cabbage cultivars. Conidia were held at each temperature for 15, 30 and 45 minutes whereas seeds were held for 30 minutes 
only (Reprinted from C. Y. Lin. 1 98 1 . Studies on downy mildew of Chinese cabbage caused by Peronospora parasitica. 
Proc. First Intern. Symposium, Asian Vegetable Research Development Centre, Shanhua, Taiwan: pp. 105-1 12. by permission 
of the author and the publisher Asian Development Research Development Centre, Taiwan). 



72 



Table 15. An analysis of sporulation of P. parasitica on cabbage cotyledons 
at two temperatures and in free water or at atmospheric water 
potentials of or -30 bars (Reprinted from H. Hartman, J.C. Sutton 
and R. Procter. 1983. Effects of atmospheric water potentials, free 
water and temperature on production and germination of 
sporangia in Peronospora parasitica. Can. J. Plant Pathol. 5:70-74, 
by permission of the authors and the publisher the Canadian 
Phytopathological Society) 



Numbers of sporangia/45 mm2 cotyledon* and integers** assigned 
for contrasts of these numbers in the following treatments!: 



Contrast 


13°C 


13°C 


13°C 


18°C 


18°C 


18°C 


F 


number 


FW 


i|/=0 


\|/=-30 


FW 


i|/=0 


\|/=-30 


test$ 




1978§ 


1466 


2265 


1042 


821 


855 




1 


-1 


-1 


-1 


1 


1 


1 


P=0.01 


2 





-1 





1 








NS 


3 











1 


-1 





NS 


4 





-1 


1 











P=0.01 


5 














-1 


1 


NS 


6 


-1 


1 














P=0.05 


7 


-1 





1 











NS 



Numbers of sporangia were contrasted using nonorthogonal coefficients and 

the differences were assessed by an F test. F tests are approximate for 

nonorthogonal contrasts. 
** Treatments assigned positive integers were contrasted with those assigned 

negative integers. 
t Treatments are identified according to temperatures and free water (FW) or 

atmospheric water potentials (i|/ in - bars) at germination. 
§ This value is the summation of mean numbers of sporangia/mm2 for each of 

45 pairs of cotyledons. 
$ Significance level or nonsignificance (NS) for results of the F test. 



73 

Localized infection also occurs in young tissues, especially those still in active 
division. In the hypertrophy caused by Albugo (Awasthi et al., 1997), the cells of the 
epidermis and cortex are dividing and may readily give entrance to Peronospora. Young 
inflorescence may wholly or partly be infected while normal tissues of older stems and 
leaves below the initial site of infection may remain free. 

When environmental conditions are suitable, conidia of P. parasitica on the surface 
of a susceptible host form germ tubes from which appressoria develop (Preece et al., 
1967). In cauliflower, appressoria are found at the junction of the anticlinal walls of the 
epidermal cells. The contents of the conidium pass into the appressorium from which 
an infection hypha develops (Chou, 1970). Penetration is usually direct and only 
occasionally through a stoma(Shiraishi et al., 1975). It breaks a hole, 4-5 /u in dia., 
through the cuticle and after entering the host the hypha expands to a diameter of 7-8 
(jl. The fungus grows initially in the region of the middle lamella between the anticlinal 
walls of the epidermal cells. Penetration between adjacent epidermal cells rather than 
via stomata had been earlier reported by Chu (1935). The infection hypha continues its 
growth between the cells of the host tissues branching in all directions and varying in 
diameter and form according to the size and shape of the intercellular spaces (Chou, 
1970). 

A single conidium of P. parasitica is sufficient to infect kohlrabi (Krober, 1969) and 
radish (Achar, 1992a). The disease intensity and rate of infection increases as the 
number of spores in the inoculum increases. Commensurately, more conidia are 
required to produce a comparable responses in older plants which are less susceptible 
to infection than young plants (Krober, 1969). Disease intensity increases with 
increasing inoculum concentrations up to 30,000 conidia/ml of water but further 
increases in inoculum have no significant effect on the host (Achar, 1992a). 

The rate of spore germination and host penetration are markedly affected by 
temperature. At 15°C, conidia germinate in 4-6h, appressoria form in 12h, and 
penetration occurs in 18-24h (Chu, 1935). Felton and Walker (1946), however, reported 
that on cabbage, germination of conidia and the subsequent penetration of the host 
takes place most rapidly at 8-12°C and 16°C, respectively. Jonsson (1966) found that 
development of the disease on winter rape is also favoured by temperatures of 8-16°C. 
In contrast Chou (1970) noted that at 20-25°C, infection occurs within 6h of the 
conidium deposition on the surface of the host cotyledon. 

Haustoria develop from the infection hyphae in the epidermis as well as in those 
of the inner tissues (Chu, 1935). The typical symptoms of infection by P. parasitica 
begin to appear two days after inoculation at 15°C, and a day or two later the formation 
of conidiophores and conidia is initiated. The haustoria in turnip and radish roots are 
at first spherical to pyriforms, becoming cylindrical or clavate, often di- or 
trichotomously branched; the maximum dimensions of an unbranched haustorium in 



74 

this situation being 18 x 25 /mi compared with only 11x8 //m in the leaves. They are 
usually spherical and bi- to trilobate and 57 x 14 /mi in the stem of B. chenensis, where 
they are cylindrical or clavate and sometimes dichotomous. Some haustoria are 
surrounded by a sheath of variable extent from a collar round the neck to a third or half 
the length of the organ itself. The few full grown haustoria found completely enveloped 
in vigourous roots inoculated with the fungus are probably incapable of functioning 
(Fig. 36). 

The systemic invasion of the hypocotyls and cotyledons of cabbage seedlings may 
take place from the soil contaminated with oospores (LeBeau, 1945). Further spread 
of the pathogen is by dissemination of conidia released from conidiophores formed on 
the cotyledons and hypocotyls (Chang et al., 1963). 

The pathogen can also enter directly through the inflorescence axis (Jang and 
Safeeulla, 1990d). The infection through the stigma and ovary wall results in 
embryonal infection. 

Pathogenesis in a susceptible combination is accompanied by large increases in 
electrolyte leakage, and increased activity of the enzymes, B-glucosidase, ribonuclease 
and peroxidase (Kluczewski and Lucas, 1982). The large increase in B-glucosidase 
originates from the pathogen and the enhanced ribonuclease activity is due to a new 
post infectional form of the enzyme. Infected B.juncea produce cellulase, indo-PMG and 
endo-PG (Singh et al., 1980). 



5. DISEASE CYCLE 

Downy mildew of crucifers is essentially a disease of foliar and other aerial plant 
tissues. The fungus survives as oospores in A. Candida - induced malformed 
inflorescence and senesced host tissues, as conidia on leaves and inflorescence, and as 
latent systemic mycelium in seeds or infected plant debris. Infections are favoured by 
temperatures between 10 and 15°C and by high atmospheric humidity following rain 
or heavy dew. The conidia produce germ tubes which penetrate anticlinal cell walls 
often on the lower surface of the leaves. The penetration is usually direct but 
occasionally also occurs through a stoma (Shiraishi et al., 1975). Primary infection from 
soil-borne oospores has been obtained (LeBeau, 1945; Chang et al., 1963). Transmission 
by infected seed is possible but its importance has not been well documented. Further 
spread of the pathogen is by dissemination of conidia released from conidiophores 
formed on the cotyledons or hypocotyls. The true leaves are usually infected through 
wind-borne conidia, resulting in spread of the disease through secondary infection. The 
pathogen dispersal over short distances in water droplets can also occur. Although, 
there is no exact information on the relationship between leaf and floral infection under 



75 




Fig. 36. Growth of crucifer downy mildew, Peronospora parasitica, in cotyledon tissues of Brassica. (A) intercellular hyphae forming 
club-shaped intracellular haustoria in host cells, stained with trypan blue (x 250); (B) fluorescence micrograph of similar 
preparation, stained with aniline blue, showing bright collars, presumed to callose-like material of host origin, at sites of 
haustorial penetration (x 280); (C) electron micrograph of intercellular hypha (I) and haustorium (H) in host cell (HC). A 
second haustorium can be seen in the same cell (x4200); and (D) cell wall encasement surrounding developing haustorium 
at site of attempted penetration. Such host cell responses are commonly seen during development of the pathogen in partially 
resistant hosts (x 10,500). (Reprinted from J. A. Lucas, J.B.R. Hayter and I.R. Crute. 1995. The downy mildews: host 
specificity and pathogenesis, IN: Pathogenesis and host specificity in plant diseases. Volume 2, Eucaryotes, Chapter 13:217- 
238, by permission of the authors and the publisher Elsevier Science ltd., The Boulevard, Langford Lane. Kidlington OX5 
1GB, U.K.). 



76 



natural conditions, most inflorescence infections, as is Albugo Candida, probably result 
from secondary spread of the pathogen rather than systemic infection. The 
diagrammatic life cycle of the disease developed by Lucas et al (1995) is given in Fig. 
37. 



6. EPIDEMIOLOGY 



In epidemics of downy mildews, the pathogen population starts from a low level of 
initial inoculum which then increases exponentially through successive cycles on the 
host during the growing season. Therefore downy mildew of crucifers is a compound 
interest disease. The seasonal increase of the pathogen population has been 
investigated much more thoroughly than that of the initial inoculum. Information has 
been generated on the multiplication phase of the disease which relates to the sequence 
of events in the life of the pathogen on its host, which are infection, colonization and 
sporulation. 

a. Disease development in relation to temperature, humidity, rainfall and 
leaf wetness 

The relationship of host-pathogen-environment interaction in case of downy 
mildew of crucifers is a complex phenomenon which determines the rate of disease 
development (Fig. 38). Among the major environmental factors which markedly 
influence the development of downy mildew are air temperature and relative humidity. 
The rates of spore germination and host penetration is affected by temperature 
variations. Chu (1935) found that at 15 °C conidia germinate in 4-6h, appressoria form 
in 12h, and penetration occurs in 18-24h. According to Eddins (1943) the downy mildew 
of cabbage is most destructive when the temperature ranges between 10° and 15°C, and 
when the plants remain wet until mid-morning for 4 consecutive days. However, Felton 
and Walker (1946) reported that on cabbage, germination of the conidia (Fig. 39) and 
subsequent penetration of the host takes place most rapidly at 8-12°C and 16°C, 
respectively. Formation of haustoria and growth of the fungus in the host tissues is 
most rapid at 20-24°C (Fig. 40). Symptoms develop quickly at 24 °C, but sporulation 
and reinfection is limited at 24 °C and 28 °C. The lower temperature of 16 °C results in 
slower growth of both the host and the pathogen, less damage, more prolific sporulation, 
more reinfection, and consequently, more profuse disease development. The severity 
of the disease at 10-15 °C seems to be the effect of temperature upon production of 
inoculum, spore germination and infection (Figs. 41, 42). However, according to 
Nashaat (1997) a temperature of 15 °C seems to be the most favourable for epidemic 
development as this favours slower growth of both host and pathogen resulting in less 
drastic damage and hence more profuse disease development. By contrast, Chou (1970) 
noted 20-25 °C, and Nakov (1972) found 15-20 °C as the most favourable temperature 



77 



o 



o 




Airborne conidia 



I 



Germinating conidium 
with appressorium and 
haustoria in epidermal cells 



* 




Oospores 



ASEXUAL 
CYCLE 



SEXUAL 
CYCLE 



♦ 





Conidiophore with conidia 



Intercellular hyphae 
forming intracellular haustoria 




Antheridium and oogonium 



Fig. 37. Diagramatic life cycle of Peronospora parasitica causing downy mildew of crucifers (Reprinted from J. A. Lucas, J.BR. 
Hayter and I.R. Crute. 1995. The downy mildews: host specificity and pathogenesis, IN: Pathogenesis and host specificity 
in plant diseases, Volume 2, Eucaryotes, Chapter 13:21 7-238, by permission of the authors and the publisher Elsevier Science 
ltd., The Boulevard, Langford Lane, Kidlington 0X5 IGB, U.K.). 



78 




Fig. 38. The relationship of host, pathogen and environment in the interaction phenotype of downy mildew of crucifers. 



79 



100 




12 16 

TEMPERATURE («C ) 



20 



2* 



Fig. 39. Effect of time and temperature on germination of conidia of Peronospora parasitica (Reprinted from M.W. Felton and J.C. 
Walker. 1946. "Environal factors affecting downy mildew of cabbage." J. Agric. Res. 72: 69-81, by permission of the 
authors and the publisher superintendent of Documents, United States Government Printing Office, Washington, DC). 



80 




12 16 

TEMPERATURE l*C) 



Fig. 40. Effect of temperature upon penetration and development of haustoria of Peronospora parasitica (Reprinted from M.W. Felton 
and J.C. Walker. 1946. "Environal factors affecting downy mildew of cabbage." J. Agric. Res. 72: 69-81, by permission 
of the authors and the publisher superintendent of Documents, United States Government Printing Office, Washington, D.C.). 



81 



132 



INITIAL SPORULATION 



— APPEARANCE OF SYMPTOMS 
HIGH HUMIDITY 



APPEARANCE OF SYMPTOMS 
LOW HUMIDITY 




4< l 



l« 



I* 23 

TEMPERATURE <"C.) 



2« 



Fig. 41 . Effect of five different temperatures on the initial spoliation of Peronospora parasitica at high humidity and upon initial 
appearance of symptoms at low and at high humidity (Reprinted from M.W. Felton and J.C. Walker. 1946. "Environal 
factors affecting downy mildew of cabbage." J. Agric. Res. 72: 69-81, by permission of the authors and the publisher 
superintendent of Documents, United States Government Printing Office, Washington, D.C.). 



82 



RELATIVE SlZ£ . 
or PLANTS 



RELATIVE SPREAD 

AND DEVELOPMENT 

Of LESIONS 



RELATIVE NUMBER 
or LESIONS 




PROPORTION 

or lesions 

SPORULATINC 



SEVERITY 
or DISEASE 



III III ill III III 

. . . . I • • 

III III III LI . . 



In III ill III -i. I., ill 

123 123 123 123 123 123 123 123 




NORMAL +N ♦*. *P -N -K 



-S 



Fig. 42. Graphic summary of infection by and development of Peronospora parasitica on cabbage plants grown in sand culture 
supplied with various nutrient solutions (Reprinted from M.W. Felton and J.C. Walker. 1946. "Environal factors affecting 
downy mildew of cabbage." J. Agric. Res. 72: 69-81, by permission of the authors and the publisher superintendent of 
Documents, United States Government Printing Office, Washington, DC.). 



83 

for infection. In temperate coastal regions of Madison, Wisconsin, USA where Chinese 
cabbage is grown from late summer through the winter and spring, downy mildew 
thrives during periods of frequent rains and high humidity. There is an 8-12h 
requirement of 100% RH for the production and dissemination of its air-borne conidia. 
Once inside the Chinese cabbage, hyphae spread through the leaves, petioles and stems, 
first feeding on the cells without apparent injury then suddenly causing yellowing, 
collapse and death of the tissues. Conidiophores and conidia are produced primarily on 
the lower side of the leaves (Williams and Leung, 1981). 

On brassica oilseeds, P. parasitica is favoured by temperatures of 8-16°C, moist 
air, and weak light (Jonsson, 1966; D'Ercole, 1975). According to Bains and Jhooty 
(1979), a 17°C and 51 mm rainfall results in low infection of mustard in contrast to high 
infection at 14 °C and 152 mm rainfall during the crop season. In a subsequent study, 
15 - 20 °C were the best temperatures for infection and development of downy mildew. 
At this temperature regime infection occurs within 24h of inoculation (Table 16, Fig. 
43). The infection frequency is reduced at 25°C with no infection observed at 30°C 
(Table 16, Fig. 43). The maximum area under disease progress curve occurs at 20 °C 
(AUDPC-60.54%, Fig. 43). Leaf wetness duration of 4-6h at 20°C, and for 6-8h at 15 °C 
is essential for severe infection and disease development on mustard (Tables 17, 18, 
Figs. 44, 45). The infection frequency and disease development increases significantly 
with the increase in duration of leaf wetness (Mehta et al., 1995). According to Kolte 
et al. (1986), sunshine has a significant negative correlation, whereas total rainfall has 
a significant positive correlation with A. Candida - induced staghead development on 
rapeseed-mustard (Table 19, Fig. 46). A reduced period of sunlight (2-6 h/d) and rainfall 
of up to 161 mm during the flowering period favours severe occurrence of the stagheads. 

In Ukraine and Russia, downy mildew of white cabbage is more severe with 
abundant rain (75 - 100 mm/10 yr) and a 14-15 h of day light (Vladimirskaya et al., 
1975). 

b. Disease development in relation to planting time 

In India, infection of mustard foliage starts by the end of October (cotyledon stage) 
and progresses up to November (Tables 20, 21). The crop planted after mid-November 
may not contract downy mildew. However, downy mildew growth as a mixed infection 
with white rust on floral parts can be seen up to March (Saharan, 1984; Kolte et al., 
1986; Mehta, 1993). 

c. Disease development in relation to host nutrition 

Peronospora parasitica is severe on cauliflower plants which suffer from potash 
deficiency, while plants with a sufficient quantity of potash are only slightly attacked 



84 



100 r 



D 

I 
S 

E 
A 
S 

E 

I 

N 
C 

I 

D 

E 
N 
C 

E 

L 
O 
G 



10 




1 

24 48 72 96 

PER CENT DISEASE INCIDENCE RECORDED AFTER INOCULATION (h) 
— 10 C — f- 15 C -*- 26 C 

-e- 25 c -*- 30 c 



F.g. 43. Progression of downy mildew (Peronospora parasitica) of mustard (Brassicajuncea) in relation to temperature (AUDPC) 
(Reprinted from N. Mehta, G.S. Saharan and O.P. Sharma. 1 995. Influence of temperature and free moisture on the infection 
and development of downy mildew of mustard. Plant Dis. Res. 10:1 14-121, by permission of the authors and publisher) 



85 



100 



s 

E 
A 
S 

E 

I 

N 
C 
I 

D 
E 
N 
C 
E 

L 
O 
G 



,P^zt— g^#=^^^ 








6 12 18 

LEAF WETNESS DURATION (H) 



24 



■^24 h 



48 h -fc- 72 h -B- 96 h 



Fig. 44. Effect of leaf wetness duration on the development of downy mildew {Peronospora parasitica) infection on mustard (Brassica 
>/icea) cultivar RH-30 at 20°C (Reprinted from N. Mehta, G.S. Saharan and OP. Sharma. 1995. Influence of temperature 
and free moisture on the infection and development of downy mildew of mustard. Plant Dis. Res. 1 0:1 14-121, by permission 
of the authors and publisher). 



86 



100 r 



D 

I 

S 
E 
A 
S 
E 

I 

N 
C 

I 

D 
E 
N 
C 
E 

L 
O 
G 




6 12 18 

LEAF WETNESS DURATION (H) 



24 



24 h 



48 h -*- 72 h -B- 96 h 



Fig. 45. Effect of leaf wetness duration on the development of downy mildew (Peronospora parasitica) on mustard (Brassica juncea) 
seedlings of cultivar RH-30 at 15°C (Reprinted from N. Mehta, G.S. Saharan and OP. Sharma. 1995. Influence of 
temperature and free moisture on the infection and development of downy mildew of mustard. Plant Dis. Res. 10:1 14-121, 
by permission of the authors and publisher). 



87 




5 too 

^ 90 



20 



RAINFALL 
Q RAINY DAYS 



I 




*a 










35 

50 



□ 



RELATIVE HUMIDITY 



,__.*/* TEMP 




y. 



r, 



Vs 



* >s 



r. 



Yr 



Y n 




Fig. 46. Weather factors associated with occurrence (A) and no occurrence (B) periods of staghead phase of white rust (Albugo 
Candida) and downy mildew (Peronospora parasitica) on mustard (Brassica juncea) in crop seasons Y, (1976-77), Y 2 (1977- 
78), Y 3 ( 1 978-79), Y 4 ( 1 979-80), Y, ( 1 980-8 1 ), Y 6 ( 1 98 1 -82) and Y, ( 1 982-83). Symbol Y. represents the number of crop 
seasons covering the period from 1977-78 through 1982-83 under no occurrence periods of stagheads (B) (Reprinted from 
S.J. Kolte, R.P. Awasthi and Vishwanath. 1986. "Effect of planting dates and associated weather factors on staghead phase 
of white rust and downy mildew of rapeseed and mustard." Indian J. Mycol. Plant Pathol. 16: 94-102, by permission of the 
authors and the society of mycology and Plant Pathology, Udaipur. India). 



88 

(Quanjer, 1928). Cabbage plants grown in soil fertilized with less potash and more 
phosphorus are more prone to downy mildew than cabbages grown in unfertilized soil 
(Townsend, 1935). However, according to Butler and Jones (1949) there is no consistent 
effect of fertilizers on the development of downy mildew of brassicas. Felton and 
Walker (1946) found no direct relationship between mildew incidence and any excess 
or deficiency of nitrogen, phosphorus or potash. On radishes tubers, conidiophores and 
conidia appear to be relatively large which is probably due to the availability of ample 
nutrient supply in the tubers (Hammarlund, 1931). 



d. Disease interaction with insecticidal sprays 

The incidence of downy mildew in plots of broccoli sprayed with emulsifiable 
insecticide formulations containing a solvent and a wetting agent is significantly 
greater than in plots sprayed with an insecticide formulation containing no solvent or 
wetting agent or in unsprayed plots (Natti et al., 1956). It is possible that emulsifiable 
insecticide formulations remove the bloom from the leaves and dissolve the wax from 
the cuticle of the leaves creating conditions favourable for the germination of P. 
parasitica spores (Natti et al., 1956). 



7. MIXED INFECTION AND ASSOCIATION WITH WHITE RUST 

The association of downy mildew and white rust infection on oilseed brassicas, 
vegetable brassicas, wallflowers and stocks have long been observed (Butler, 1918; 
Wiese, 1927). 

On horse-radish leaves and petioles, A. Candida (white rust) and P. parasitica 
(downy mildew) are frequently associated with each other causing brown rot 
commencing at the head of the rootstock and extending downwards (Boning, 1936). 

According to Bains and Jhooty (1985), A. Candida often appears first in combined 
infections. It is followed by infection with P. parasitica which develops in and around 
A. Candida colonies (Fig. 3). A. Candida predisposes the host tissues towards 
susceptibility to this pathogen. The development of hypertrophied tissues of the 
staghead phase are attributed to infection with A. Candida. The intensity of mixed 
infection by A. Candida and P. parasitica on B. juncea inflorescence has been reported 
to be from 0.5 to 29.0 percent under Punjab (India) conditions (Bains and Jhooty, 1979, 
1985). There is preferential parasitism of P. parasitica on galls of B. campestris caused 
by A Candida (Chaurasia et al., 1982). The hypertrophied malformed floral organs of 
mustard infected with A. Candida are usually heavily covered with white sporulating 
fungal growth of P. parasitica consisting of conidia and conidiophores (Saharan and 



89 



Table 16. Effect of temperature on infection by Peronospora parasitica 
and disease development on mustard seedlings (cv. RH-30) (Mehta, 
Saharan and Sharma, 1995) 



Temp 




Per cent Disease Incidence after inoculation (h) 










(°C) 






















< 


24 


48 


72 


96 


Mean 


AUDPC* 


10 


0.00 


(1.81) 


54.09 (47.52) 


64.19 (53.52) 


70.53 


(54.49) 


47.20 


(40.08) 


43.83 


15 


2.91 


(6.71) 


58.47 (50.30) 


73.88 (60.37) 


75.55 


(62.44) 


52.70 


(44.88) 


45.86 


20 


34.31 


(35.59) 


78.33 (62.49) 


87.56 (69.98) 


90.27 


(75.00) 


72.61 


(60.76) 


60.54 


25 


0.00 


(1.81) 


5.08 (9.32) 


14.25 (20.90) 


19.72 


(25.82) 


9.76 


(14.26) 


14.89 


30 


0.00 


(1.81) 


0.00 (1.81) 


0.00 (1.81) 


0.00 


(1.81) 


0.00 


(1.81) 


1.81 


Mean 


9.40 


(9.54) 


48.99 (34.23) 


59.97 (41.51) 


64.01 


(44.51) 




- 


- 


Correlation 


















coefficient 


0.08 


0.78 


0.82 




0.80 








(r) 





















LSD (0.05) Temperature (T) 3.55 Observations (0) 3.55 Temp, x Observation : 7. 11 

(TX0) 

Figures in the parentheses are angular transformed values after adding 0.1. 
* Area under disease progress curve 



Verma, 1992; Saharan, 1992a). Incidence and severity of mixed infections by A. 
Candida and P. parasitica on B. juncea inflorescence is higher on detopped than on 
normal plants (Bains, 1989). Severity of mixed infections on leaves is not related to 
infections on inflorescence. It seems that greater susceptibility of new inflorescence 
and their availability over extended periods of time is associated with this pheno- 
menon (Bains and Sokhi, 1986). Peronospora parasitica also causes severe infections 
and high levels of sporulation on plants of mustard systematically infected with 
mustard mosaic virus (Bains and Jhooty, 1978). 

In artificially inoculated leaves of mustard, the stimulatory effect of A. Candida 
infection is more intense when P. parasitica is inoculated 7 days after A. Candida 
(Chaudhury and Verma, 1987). When Peronospora and/or Albugo are inoculated alone 
or in different combinations, the downy mildew infection takes 7 days, while white rust 
appears within 5-6 days of inoculations. When both the pathogens are inoculated 
simultaneously in a 50:50 spore concentration then there is delay in the expression of 
infections by Peronospora for 2-3 days (Mehta et al., 1995). 



90 



Table 17. Effect of leaf wetness duration on infection by Peronospora 
parasitica and disease development on mustard seedlings (cv. RH- 
30) at 20°C (Mehta, Saharan and Sharma, 1995) 



Leaf Wetnes 


Percent Disease Incidence after inoculation (h) 










Duration (h) 


24 




4 


18 


72 


96 


Mean 





0.0 


(1.81) 


0.00 


(1.81) 


0.0 


(1.81) 


7.08 


(15.41) 


1.77 


(5.21) 


2 


0.0 


(1.81) 


0.00 


(1.81) 


5.18 


(13.30) 


8.51 


(16.57) 


3.42 


(8.37) 


4 


0.0 


(1.81) 


3.05 


(8.94) 


7.77 


(16.14) 


29.04 


(32.48) 


9.96 


(14.84) 


6 


0.0 


(1.81) 


6.80 


(15.05) 


22.12 


(27.87) 


29.35 


(32.63) 


14.56 


(19.34) 


8 


0.0 


(1.81) 


21.52 


(27.00) 


35.83 


(35.99) 


35.30 


(38.69) 


24.16 


(25.79) 


10 


0.0 


(1.81) 


24.58 


(29.70) 


45.37 


(42.39) 


44.44 


(41.85) 


28.59 


(28.93) 


12 


0.0 


(1.81) 


18.57 


(25.30) 


53.03 


(46.84) 


62.26 


(52.24) 


33.46 


(34.05) 


14 


0.0 


(1.81) 


25.27 


(30.18) 


56.46 


(48.78) 


61.94 


(52.02) 


35.91 


(33.19) 


16 


0.0 


(1.81) 


27.91 


(31.79) 


56.52 


(48.84) 


62.50 


(52.31) 


36.73 


(33.68) 


18 


5.0 


(8.83) 


26.80 


(31.22) 


59.86 


(50.78) 


68.33 


(55.88) 


39.99 


(36.68) 


20 


9.16 


(15.15) 


34.38 


(35.48) 


61.57 


(51.80) 


68.61 


(56.06) 


43.43 


(39.62) 


22 


26.25 


(30.39) 


46.11 


(42.82) 


61.66 


(57.98) 


69.16 


(56.04) 


50.79 


(45.30) 


24 


32.96 


(35.01) 


53.19 


(46.90) 


67.80 


(55.96) 


72.77 


(58.83) 


56.68 


(49.17) 


Mean 


5.64 


(8.12) 


22.16 


(25.23) 


41.01 


(37.88) 


47.94 


(43.13) 


29.18 


(28.59) 


LSD (0.05) 


Observation (0) 


= (2.54) 


Leaf Wetness duration (W) = (4.59) 


0*W = (9.19) 


Correlation 






















coefficient (r) 




0.72 


0.95 


0.95 




0.94 









Figures in parentheses are angular transformed values after adding 0.1. 



Histopathological studies carried out by Mehta et al. (1995) indicated that conidia 
of Peronospora and sporangia of Albugo inoculated on mustard leaves germinate 24h 
after inoculation. Two days after inoculation infection is normally confined to the host 
epidermis. The pathogens penetrate up to 1/3 of the mesophyll cells by the third day 
after inoculations. Six days after inoculation, the pathogens progress deeper into the 
tissues. When P. parasitica is inoculated prior or after A. Candida, mycelium can be 
seen in the intercellular spaces with globose to knob like haustoria in the mesophyll 
cells. When A. Candida is inoculated alone or in combination with P. parasitica, the 
pathogen emerges from the lower epidermis and forms pustules. However, on its own 
Peronospora causes necrosis in the mesophyll cells. When both the pathogens are 
inoculated together, the infection is confined to the upper layer of the mesophyll with 
limited colonization of the cells and few haustoria or mycelium in the intercellular 
spaces. Nine days after inoculation, characteristic disease symptoms are visible. The 
white rust pustules show hyaline sporangiophore bearing globose to oval shaped 



91 



Table 18. Effect of leaf wetness duration on infection by Peronospora 
parasitica and disease development on mustard seedlings (cv. RH- 
30) at 15°C (Mehta, Saharan and Sharma, 1995) 



Leaf Wetness 




Percent Disease Incidence after inoculation (h) 








Duration (h) 




24 




48 


i 


72 


j 


?6 


Mean 





0.0 


(1.81) 


0.00 


(1.81) 


0.00 


(1.81) 


0.00 


(1.81) 


0.00 


(1.81) 


2 


0.0 


(1.81) 


0.00 


(1.81) 


0.00 


(1.81) 


3.75 


(9.84) 


0.93 


(3.81) 


4 


0.0 


(1.81) 


0.00 


(1.81) 


0.00 


(1.81) 


10.00 


(18.21) 


2.50 


(5.91) 


6 


0.0 


(1.81) 


1.66 


(5.55) 


1.66 


(5.55) 


20.83 


(26.90) 


8.52 


(9.95) 


8 


0.0 


(1.81) 


34.02 


(35.73) 


47.10 


(43.41) 


47.10 


(43.41) 


32.05 


(31.09) 


10 


0.0 


(1.81) 


49.32 


(44.69) 


56.41 


(48.74) 


56.41 


(48.74) 


40.53 


(35.99) 


12 


0.0 


(1.81) 


46.84 


(43.26) 


58.05 


(49.74) 


58.47 


(49.99) 


40.84 


(36.20) 


14 


0.0 


(1.81) 


53.82 


(47.18) 


62.73 


(52.36) 


62.78 


(52.48) 


44.83 


(38.00) 


16 


0.0 


(1.81) 


54.02 


(47.39) 


63.31 


(52.81) 


63.31 


(52.81) 


45.16 


(38.70) 


18 


0.0 


(1.81) 


57.93 


(49.55) 


64.60 


(53.49) 


66.97 


(54.94) 


47.37 


(39.49) 


20 


0.0 


(1.81) 


65.83 


(54.31) 


67.50 


(55.33) 


67.50 


(55.33) 


50.20 


(41.69) 


22 


0.0 


(1.81) 


67.50 


(55.33) 


68.09 


(55.69) 


68.88 


(56.17) 


51.11 


(42.25) 


24 


0.0 


(1.81) 


70.07 


(57.06) 


70.07 


(57.06) 


70.07 


(57.06) 


52.55 


(46.51) 


Mean 


0.0 


(1.81) 


38.53 


(34.28) 


43.04 


(38.53) 


45.85 


(42.26) 


31.85 


(29.22) 


LSD (0.05) 


Observation (0) = 


= (1.38) 


Leaf Wetness duration (W) = (2.49) 


0*W 


= (4.99) 


Correlation 






















coefficient (r) 


i 


0.00 


0.91 


0.91 




0.93 









Figures in parentheses are angular transformed values after adding 0.1. 



sporangia in chains. The Peronospora mycelium is intercellular with lobe shaped 
haustoria in the distorted tissue of leaves. When both pathogens are inoculated 
together, infection is extended to mesophyll cells and there is development of pustule 
below the epidermis. Twelve days after inoculation, complete colonization of the host 
tissues by the pathogen is evident from the development of necrotic zone by P. 
parasitica and bursting of pustules releasing sporangia in case of A. Candida. In the 
inflorescence, the mycelium passes through the epidermis, hypodermis, cortex and 
finally reaches to the pith region. The mycelium is in abundance in the cortex and 
produces conidiophores bearing conidia above the epidermis layer. For A. Candida, 
numerous sporangiophores bearing sporangia are observed below the epidermis layer 
in the form of pustules and knob like haustoria in the tissues. In the colonized tissues 
both pathogens cannot be distinguished based on somatic morphology. 



92 

Table 19. Prediction equations for the progress of downy mildew and white 
rust of rapeseed-mustard using different combinations of weather 
factors (Reprinted from S.J. Kolte, R.P. Awasthi and Vishwanath. 
1986. Effect of planting dates and associated weather factors on 
staghead phase of white rust and downy mildew of rapeseed and 
Indian J. Mycol. Plant Pathol. 16:94-102, by permission of the 
authors and the publisher the Society of Mycology and Plant 
Pathology, Udaipur, India) 



Equations 



x 6 



R 



Staghead incidence +16.925 +0.019 -0.132 -0.086 +0.158 +0.030-1.469 0.6849 

(Yj) (%) 
Staghead severity +86.169 -1.241 -0.129 -0.503 +0.054 +0.472-2.125 0.6283 

(Y 2 ) (%) 



x x = mean maximum temperature 
x 3 = mean relative humidity 
x 5 = total rainy days 
* Significant at 5% level 



x 2 = mean minimum temperature 

x 4 = total rainfall (mm) 

x 6 = mean bright sunshine period (h/day) 



8. BIOCHEMISTRY OF THE HOST PATHOGEN INTERACTION 

Biochemical studies of the growth and survival of a pathogen and the changes it 
induces in its host can ultimately lead to a better understanding of the disease 
development, its epidemiology and control. Ideal prerequisites for meaningful studies 
of the biochemistry of the host-parasite interaction are: (a) A clear understanding of the 
genetic control of virulence and avirulence in the parasite and of susceptibility and 
resistance in the host, (b) Precise histological and cytological descriptions of spore 
germination, infection, and the establishment and development of the host-pathogen 
interaction, and (c) The availability of methods for maintaining the pathogen alone and 
in combinations with its host, under controlled conditions. Unfortunately, these criteria 
have not yet been satisfactorily met for downy mildew of crucifers. 



a. Metabolic changes 

Many marked shifts were observed in the metabolic processes of plant tissues 
following infection by biotrophic parasites. These included changes in respiration, 
photosynthesis, nucleic acid and protein synthesis, and phenol metabolism. There could 



93 



Table 20. Effect of planting time on the severity of downy mildew and white 
rust of mustard (Saharan, 1984) 



Planting time 



Hisar 



Percent disease intensity 



Kanpur 



Pantnagar 



06.10.1978 


10.0 


21.10.1978 


8.6 


28.10.1978 


18.6 


06.11.1978 


55.4 


18.11.1978 


68.5 


02.12.1978 


72.8 


01.10.1979 


- 


10.10.1979 


4.6 


20.10.1979 


10.0 


30.10.1979 


22.5 


09.11.1979 


46.8 


20.11.1979 


57.5 


03.10.1981 


- 


23.10.1981 


- 


13.11.1981 


_ 



24.16 


- 


28.30 


- 


34.34 


- 


36.18 


- 


40.91 


- 


46.15 


- 


- 


15.04 


- 


19.85 


- 


32.85 



also be changes in the translocation and accumulation of nutrients and in the levels of 
endogenous growth substances. 

The respiration rate was raised sharply soon after P. parasitica infection of 
cabbage cotyledons and reached a maximum, almost twice that of uninfected controls, 
at the time of the initiation of sporulation (Fig. 47). The chlorophyll content of infected 
and noninfected cotyledons did not differ significantly at any time (Fig. 48). The 
increased respiration rate of the infected tissues did not reflect any significant changes 
to the pentose phosphate pathway in this infection since no acyclic polyhydric alcohols 
were detected in soluble extracts of either infected leaves or fungal conidida (Thornton 
and Cooke, 1974) (Figs. 49, 50). 

Pathogenesis in Brassica - Peronospora combinations was observed to be 
accompanied by large increases in electrolyte leakage (Fig. 54-57) and increased activity 
of /?.-glucosidase (Fig. 55), ribonuclease (Fig. 56) and peroxidase (Fig. 57) (Kluczewski 
and Lucas, 1982). The large increase in /?.-glucosidase were of pathogen origin while 
enhanced ribonuclease activity was due to a new post-infectional form of the enzyme. 



94 



4» 
Q. 



0~ » 

o I 

• eg 

o O 



200 - 



150 - 



(C f> 




100 - 



2 3 4 

Days after inoculation 



Fig. 47. Rates of 2 uptake of infected and uninfected cotyledons at various times after inoculation: (•-•), infected; (o-o), uninfected; 
A, visible signs of sportulation (Reprinted from J.D. Thornton and R.C. Cooke. 1974. Changes in respiration, chlorophyll 
content and soluble carbohydrates of detached cabbage cotyledons following infection with Peronospora parasitica (Pers. 
ex FY.), by permission of the authors and the publisher Academic Press Limited, London). 



95 



Chlorophyll„+ Chlorophyll,, 
(mg x I0" 2 per g fresh wt) 





X* <J> CD 

o o o 


o 
o 




1 1 1 


1 

• o- 


r 




.^ 


ro 


— 


• o - 


o 






o 






v> CM 


c^^ 




o 

-* 
(0 


A 




2. * 


• o 


— 


5" 
o 
o 

c 


// 




o m 


• o 


— 


o 

3 


// 




0> 


•o 

IS 




-J 


1 1 1 


1 



Fig. 48. Chlorophyll a plus chlorophyll,, content of infected and uninfected cotyledons at various times after inoculation: (•-•), infected; 
(o-o), uninfected (Reprinted from J.D. Thornton and R.C. Cooke. 1974. Changes in respiration, chlorophyll content and 
soluble carbohydrates of detached cabbage cotyledons following infection with Peronospora parasitica (Pers. ex Fr), by 
permission of the authors and the publisher Academic Press Limited, London). 



96 











glucose 






2 












- 


10 






[ P 


r 


i 


r 


r 


- 



£ C 2 

• S io 

o *- 

•5 • 

■fi JJ 



o 
u 



1 


n 


JC 


»os 


e 


































- 



I0h sucrose 
5 



n rH m J ffl jj rfl I 



10 h Inositol 
5 



^ ^ jt m Tn Th n tr 



I 2 3 4 5 6 7 
Days after inoculation 



8 



Fig. 49. Carbohydrate content of the alcohol soluble fraction of infected and uninfected cotyledons at various times after inoculation 
with Peronospora parasitica: U = infected; □ = uninfected; T = trace (indicating that the peak height of the TMS derivative 
was indeterminable at an attenuation of 20 x IO 3 (Reprinted from J.D. Thornton and R.C. Cooke. 1974. Changes in 
respiration, chlorophyll content and soluble carbohydrates of detached cabbage cotyledons following infection with 
Peronospora parasitica (Pers. ex Fr.), by permission of the authors and the publisher Academic Press Limited, London). 



97 



CO 



O 
O 



». >» 

Tf — 

>» o 

jz o 



o 
o 



o. 

E 



004 












- 


03 


- 










- 


02 
















- 


01 
















"1 






n-n 


- 



glucose fructose trehalose 

sucrose inositol 



Fig. 50. Principal carbohydrates of the alcohol soluble fraction of sporangia from infected cotyledons and control washings, 7 days 
after inoculation with Peronospora parasitica: ■ = infected; □ = uninfected (Reprinted from J.D. Thornton and R.C. Cooke. 
1974. Changes in respiration, chlorophyll content and soluble carbohydrates of detached cabbage cotyledons following 
infection with Peronospora parasitica (Pers. ex Fr.), by permission of the authors and the publisher Academic Press Limited, 
London). 



98 

Table 21. Influence of planting dates on staghead incidence and severity of 
white rust and downy mildew of rapeseed and mustard in three rabi 
crop seasons starting from 1977-78 to 1979-80 (Reprinted from S.J. 
Kolte, R.P. Awasthi and Vishwanath. 1986. Effect of planting dates 
and associated weather factors on staghead phase of white rust and 
downy mildew of rapeseed and mustard. Indian J. Mycol. Plant 
Pathol. 16:94-102, by permission of the authors and the publisher the 
Society of Mycology and Plant Pathology, Udaipur, India) 







1977-78 




1978-79 






1979-80 








Yellow 




Yellow 






Yellow 






Mustard Sarson 


Toria 


Mustard Sarson 


Toria 


Mustard Sarson 


Toria 




I S 


I S 


I S 


ISIS 


I S 


I S 


I S 


I S 


Oct. 1-6 


2.7 11.0 


0.0 0.0 


0.0 0.0 


20.6 15.6 24.3 23.1 


10.2 20.3 


8.9 20.3 


4.8 15.7 


0.7 4.4 


Oct. 11-14 


6.7 15.6 


1.2 8.4 


0.0 0.0 


14.6 20.4 23.7 24.0 


7.7 21.8 


14.2 17.7 


8.3 32.4 


1.8 6.4 


Oct. 20-22 


10.2 23.4 


4.1 24.8 


0.0 0.0 


10.6 11.3 17.2 14.9 


18.2 16.8 


7.8 14.6 


3.6 17.7 


3.6 16.9 


Oct. 31-Nov.l 


10.2 20.9 


8.2 35.2 


10.2 23.9 


7.8 9.7 11.6 17.2 


24.9 13.5 


5.3 11.5 


11.2 43.4 


9.2 22.4 


Nov. 1-11 


9.3 22.4 


9.7 25.8 


10.6 25.3 


9.6 11.1 11.2 16.5 


9.4 14.4 


3.1 5.2 


6.7 16.3 


182347Nov. 


20-22 


14.2 22.8 


38.2 25.7 


22.2 24.0 


9.1 10.9 13.7 19.0 


8.9 15.3 


4.3 8.5 


1.4 8.8 


15.9 23.2 



I = Incidence (% plants affected). 
CD at 5% 



S = Severity (% recemes affected/plant) 



For planting dates 



I 
S 
I 

s 



1977-78 
5.6 
5.5 
NS 
3.4 



1978-79 
2.8 
1.7 
1.8 
1.3 



1979-80 
3.6 
9.8 
2.6 

4.2 



In vivo infected leaves of B.juncea produced cellulase, endo - PMG and endo - PG (Singh 
et al., 1980). 

b. Role of natural biochemical compounds 



There are a number of natural biochemical compounds present in host tissues 
which may influence the defence mechanism of crucifers against downy mildew 
infection. The role of phenolic compounds, glucosinolates and flavour volatile 
compounds in providing resistance to crucifers against downy mildew infection has been 
explained in section "Biochemical basis of resistance" of chapter 9E. 



99 

9. RESISTANCE 

Genetic resistance is the most important attribute of the host defense against P. 
parasitica. Host resistance provides an economical, environmentally benign, and widely 
accepted method of managing downy mildew of crucifers. 

a. Mechanism of host resistance 

The first defense barrier in crucifers is the cuticle which is often covered with a 
waxy layer, a hairy surface and a few stomata with narrow apertures. The mechanisms 
of resistance to P. parasitica in Chinese rape, cabbage and radish was studied by Wang 
(1949) through observations on pathogen entry, mycelial and haustorial development 
and sporulation. All plants regardless to whether they are susceptible or resistant, 
were initially penetrated directly through the epidermal cells or by entering the 
stomata. After penetration, the mycelia grew through the intercellular spaces of the 
leaf mesophyll and haustoria penetrate the cells of susceptible hosts. In the resistant 
and immune hosts, development of mycelia and formation of haustoria were curtailed 
with the death of the surrounding host cells (Fig. 58). The pathogen sporulated 
abundantly on susceptible hosts, but necrotic reaction was associated with the infection 
of the resistant hosts. On the immune hosts, few minute necrotic spots/or occasionally 
no visible symptoms were observed. Weak light and high moisture conditions may alter 
the resistant or immune reaction of the host. 

The growth of two isolates of P. parasitica obtained from cauliflower and oilseed 
rape (B. napus) was assessed in their respective hosts of origin and also in the 
alternative combination by Kluczewski and Lucas (1982). Both isolates were capable 
of infecting either host, but there were marked contrasts in the time course and extent 
of mycelial development, the amounts of associated host cell necrosis, and eventual 
intensity of sporulation (Figs. 51-53). Oilseed rape which is partially resistant to the 
isolate from cauliflower, exhibits extensive necrosis of mesophyll cells in conjunction 
with reduced mycelial development and delayed and reduced sporulation by the 
pathogen (Figs. 51-53). The isolate from oilseed rape is virulent on both host species. 
Pathogenesis in the susceptible combination is accompanied by a large increase in 
electrolyte leakage, and increased activity of the enzymes /?-Glucosidase, ribonuclease 
and peroxidase. 

The growth of hyphae in the susceptible cultivar of Japanese radish was reported 
to be faster than that in the resistant cultivar (Ohguchi and Asada, 1991). Five days 
after inoculation, haustoria were formed in the cells of the 63rd cell layer from the 
inoculated layer in the susceptible cultivar and in the cells of 12th cell layer in the 
resistant cultivar. The haustoria formed in both cultivars were similar in size and 
shape. On the surface of haustoria spherical or semi-spherical granules, 1.7 - 3.7 //m 



100 



12 



10 



(a) Cauliflower 



* A 



I 



AjL 



Li 



i 






>i 



>. 2 

S 



(b) Oilseed rape 



r5 



rfl 



IL 




2 3 4 5 

Days after inoculation 



3 



2 o 



Fig. 51. Relationship between mycelial development and host-cell necrosis estimated as granulation and browning of cells in (a) 
cauliflower; and (b) oilseed rape inoculated with Peronospora parasitica isolates from cauliflower (CI) and oilseed rape (Rl ). 
D CI mycelial growth index; ■ CI necrotic cell index; □ Rl necrotic cell index. Bars indicate + standard deviation 
(Reprinted from S.M. Kluczewski and J. A. Lucas. 1982. Development and physiology of infection by the downy mildew 
fungus Peronospora parasitica (Pers. ex Fr.) Fr. in susceptible and resistant Brassica species. Plant Pathology 31:373-389, 
by permission of the authors and the publisher Blackwell Science ltd., Osney Mead, Oxford, U.K.). 



101 



c 

0> 



a> 
a. 



c 
o 

o 

a. 



■o 
c 
o 
in 

3 
O 




12 3 4 

Days after inoculation 



Fig. 52. Time course of sporulation of Peronospora parasitica isolate from cauliflower (o), and oilseed rape (■) on cauliflower ( -) 
and oilseed rape (...). Bars indicate + standard deviation (Reprinted from S.M. Kluczewski and J. A. Lucas. 1982. 
Development and physiology of infection by the downy mildew fungus Peronospora parasitica (Pers. ex Fr.) Fr. in 
susceptible and resistant Brassica species. Plant Pathology 31:373-389, by permission of the authors and the publisher 
Blackwell Science ltd., Osney Mead, Oxford, U.K.). 



102 



a 






• 





•< .- - ~- „ 




Fig. 53 . Cotyledon tissue 4 days after inoculation with Peronospora parasitica cauliflower isolate stained with trypan blue and cleared 
in chloral hydrate. (A) intercellular hyphae in oilseed rape cultivar Primor showing left, developing haustoria (arrows) in host 
mesophyll cells close behind the hypha apex, and right, necrosis of penetrated host cells in older regions of a hypha. x 400; 
(b) Intercellular hyphae in cauliflower cultivar VSAG forming abundant intracellular haustoria. Note absence of host-cell 
necrosis, x 400 (Reprinted from S.M. Kluczewski and J.A. Lucas. 1982. Development and physiology of infection by the 
downy mildew fungus Peronospora parasitica (Pers. ex Fr.) Fr. in susceptible and resistant Brassica species. Plant Pathology 
31:373-389, by permission of the authors and the publisher Blackwell Science ltd., Osney Mead, Oxford, U.K.). 



103 



Oilseed rope 




12 3 4 5 

Days oft«r inoculation 



Days after inoculation 



Fig. 54. Conductivity changes of deionized glass-distilled water containing samples of uninfected cotyledons (...) and cotyledons 
infected (-) by Peronospora parasitica isolate from cauliflower (o) and oilseed rape (■). Each point represents the mean 
of four replicates (Reprinted from S.M. Kluczewski and J. A. Lucas. 1982. Development and physiology of infection by the 
downy mildew fungus Peronospora parasitica (Pers. ex Fr.) Fr. in susceptible and resistant Brassica species. Plant Pathology 
3 1 :373-389, by permission of the authors and the publisher Blackwell Science ltd., Osney Mead, Oxford, U.K.). 




I 2 3 4 5 6 

Doys after inoculation 





Oilseed rape 






60- 








50- 








40- 






| 


30- 






1 


20- 






I 


10' 












..#... 


•••• • • 



12 3 4 3 

Days after inoculation 



Fig. 55. (J-glucosidase activity in extracts of control cotyledons (...) and cotyledons infected (-) by either cauliflower (o) or oilseed 
rape (-) isolate of Peronospora parasitica (Reprinted from S.M. Kluczewski and J. A. Lucas. 1982. Development and 
physiology of infection by the downy mildew fungus Peronospora parasitica (Pers. ex Fr.) Fr. in susceptible and resistant 
Brassica species. Plant Pathology 31 :373-389, by permission of the authors and the publisher Blackwell Science ltd., Osney 
Mead, Oxford, U.K.). 



104 



?■ 



Cauliflower 




Oilseed rope 



/ 



A-^ 



I 2 3 4 5 6 

Days oMer inoculo'ion 



12 5 4? 

Ooys after moculotion 



Fig. 56. Acid ribonuclease activity in extracts of control cotyledons (...) and cotyledons infected (-) by Peronospora parasitica isolate 
from cauliflower (o) and oilseed rape (-)(Reprinted from S.M. Kluczewski and J. A. Lucas. 1982. Development and 
physiology of infection by the downy mildew fungus Peronospora parasitica (Pers. ex Fr.) Fr. in susceptible and resistant 
Brassica species. Plant Pathology 3 1 :373-389, by permission of the authors and the publisher Blackwell Science ltd., Osney 
Mead, Oxford, U.K.). 



0.8 




12 3 4 5 

Days ofte' fnocu'af'to' 1 



Oilseed rape 



r 



12 3 4 6 

['Oy< o"e' incrvilOTiO" 



Fig. 57. Peroxidase activity in extracts of control cotyledons (...) and cotyledons infected by either cauliflower (o) or oilseed rape (■) 
isolate of Peronospora parasitica (Reprinted from S.M. Kluczewski and J. A. Lucas. 1982. Development and physiology 
of infection by the downy mildew fungus Peronospora parasitica (Pers. ex Fr.) Fr. in susceptible and resistant Brassica 
species. Plant Pathology 31 :373-389, by permission of the authors and the publisher Blackwell Science ltd., Osney Mead, 
Oxford, U.K.). 



105 




Fig. 58. Peronospora parasitica brassicae rae 2. Entry of germ tube of the conidium, through (A) an epidermal cell and (B) a stoma. 
Mycelium in tissue of (C) the susceptible Chinese rape host, and (D) the immune radish host. Legend: Sp = conidium; Ap 
= appressorium; IH = infection hypha; My = mycelium; Ha = haustorium; Sh = sheath; Ep = epidermis; St = stoma; Sp = 
spongy mesophyll tissue; and DC = dead host cells (Reprinted from T.M. Wang. 1949. Studies on the mechanism of 
resistance of cruciferous plants to Peronospora Parasitica. Phytopathology 39; 541-547, by permission of the publisher 
American Phytopathological Society). 



106 

in diameter, were often observed in the susceptible cultivars, while rarely observed in 
the resistant cultivar. 

b. Host-pathogen recognition system 

The Arabidopsis - P. parasitica system was recently adopted as a model system for 
studying the recognition process for gene-for-gene interactions (Davis and 
Hammerschmidt, 1993). The determination of specificity and mechanism of recognition 
is a highly complex phenomenon. The comprehension of this phenomenon depends on 
better knowledge of molecular biology and genetics of the host-pathogen interaction. 
Cytological and biochemical studies are being attempted to identify the stages at which 
the incompatibility recognition events occur. Proposed steps between recognition, 
signal transduction and activated defence during the hypersensitive response in 
crucifers to P. parasitica are outlined in Table 22 (Lebeda and Schwinn, 1994). 

c. Systemic acquired resistance 

Systemic acquired resistance (SAR) has been demonstrated in Arabidopsis plants 
treated with chemical inducers such as 2, 6-dichloroisonicotinic acid (Uknes et al., 
1992). Resistance is expressed within a few days of exposure to the inducer and inhibits 
subsequent infection by both bacterial and fungal pathogens, including P. parasitica. 
The degree of protection varies depending upon the concentration of inducer chemical 
used, but at higher dose rate sporulation of P. parasitica is completely inhibited. 
Microscopic examination of induced plants inoculated with the fungus reveals that 
hyphal growth is restricted to the initial penetration site, associated with a necrotic 
reaction in host cells. In plants treated with lower doses of the chemical, some hyphal 
development occurs, but haustoria are reduced in size and many are encased in 
material of host origin. Host cells penetrated by haustoria often become necrotic. 
Similar cytological events occur in uninduced hosts inoculated with incompatible 
isolates of P. parasitica (Kluczewski and Lucas, 1982). Induction of SAR appears to 
enhance the efficiency of host defence responses and thereby disrupts the development 
of a biotrophic relationship in a normally compatible host. The 

biochemical mechanism of SAR is not yet understood, but models envisage a 
translocated single molecule that induces changes in tissues removed from the initial 
inoculation site. Development of SAR is associated with induction of PR proteins 
(Uknes et al., 1992). 

d. Genetics of host-pathogen relationship 

Resistance derived from the Broccoli Introduction No. PI 189028 to P. parasitica 
race 1 is found to be governed by one dominant gene. The distribution of resistant 



107 

Table 22. Expected sequence of events leading to hypersensitive reaction 
expression in crucifers to P. parasitca infection (Reprinted from 
A. Lebeda and F.J. Schwinn. 1994. The downy mildews - an 
overview of recent research progress. J. Plant diseases and 
protection 101:225-254, by permission of the authors and the 
publisher Eugen Ulmer Verlag GmbH, Germany). 

Differentiation in response to plant signal 

Production of cultivar - specific elicitors 
r Primary recognition 
Activated responses 
De novo protein synthesis in penetrated cell 
Irreversible membrane damage and release of phenolics 

Release of endogenous elicitors 
Accumulation of wall bound phenolics 
Release of secondary signals 
Secondary recognition 
Transcriptions of mRNAs controlling biosynthesis of lignin 
Precursors in surrounding 
Deposition of lignins in and around the infection site. 



plants in populations segregating for both downy mildew resistance and waxless foliage 
indicates that resistance is independent of foliage wax. Resistance to race 1 and race 
2 obtained from the cabbage introduction PI 245015 is found to be inherited 
independently. Resistance is governed by one dominant gene for each race (Natti et 
al., 1967). However, in a later study, Hoser-Krause et al. (1991) found that in broccoli, 
(B. oleracea var botrytis) resistance to a Polish isolate of P. parasitica at the 4-5 leaf 
stage is determined by a single recessive gene different from genes determining 
resistance at the cotyledon stage. Subsequently they (Hoser-Krauze et al. 1995) found 
that in broccoli resistance to downy mildew is governed by 3 or 4 dominant 
complimentary genes. Both seedling and mature plant resistance has been reported in 
B. oleracea with the later being quantitative (Dickson and Petzoldt, 1996). 

In Chinese cabbage resistance to downy mildew at the cotyledon stage expressed 
as a reduction in the sporulation capacity of P. parasitia was found to be under 
dominant monogenic control (Niu et al., 1983). However, Yuen (1991) while analysing 
Chinese cabbage lines with a reduced rate of mildew development found additive effects 
with involvement of several resistant genes. 

The resistance to P. parasitica in radish cultivars Tokinaski (All Season) and 
Okura was found to be controlled by two dominant and independent genes (Bonnet and 
Blancard, 1987). 



108 

Cytoplasmic male sterile (B. campestris) breeding lines with resistance to downy 
mildew have been identified by Leung and Williams (1983). Downy mildew resistance 
was expressed in cotyledons as a reduction in the sporulation capacity of P. parasitica. 
In high and partially resistant hosts, spore production ranged from 7 to 30 spores/g host 
tissue as compared to 260 spores/g host tissue in the fully susceptible hosts. In the 
chinensis lines, 80% of the plants showed high to partial resistance to P. parasitica 
whereas in the other lines resistance ranged between 10% to 50%. 

Differential host resistance to homologous isolates of P. parasitica has been 
identified in B. rapa (B. campestris), B. napus, B. juncea and B. oleracea. In B. napus, 
resistance in the oilseed rape cultivar Cresor is controlled by a single dominant allele 
(Lucas et al., 1988). In B. oleracea, differential resistance has been located in a land 
race cauliflower "Palermo Green". A model based on two or possible three major genes 
has been proposed by Moss et al. (1988) to account for the reaction patterns of 
individual plants to select fungal isolates within a host population (Table 23). In B. 
campestris both rapid cycling and commercial genotypes have been identified for 
differential resistance (Table 12). Four homologous isolates have identified a range of 
differential responses. 

According to Nashaat et al. (1995a, b, 1996) resistance of the RES-01-1-4 and 
RES-26 lines of B. napus to isolate P003 of P. parasitica is conditioned by a single 
dominant resistant gene, whereas resistance of RES-02 is conditioned by two 
independent dominant resistance genes. Later, Nashaat and Awasthi (1995) selected 
differential putative homozygous resistant lines from seedling populations of accessions 
that exhibited a heterogeneous reaction to the isolates from B. juncea (Tables 24, 25). 

In inoculation tests on Arabidopsis thaliana with seven pathogen isolates and 
eleven host accessions, a range of interaction phenotypes were observed including 
localized necrosis (flecking), more extensive cell collapse (pitting), delayed sporulation, 
or complete susceptibility (Holub et al., 1994). Segregation for these phenotypes among 
F 2 progeny from a half dialle cross between nine A. thaliana accessions, and ten host 
loci (termed RPP, recognition of P. parasitica loci) has been observed. Four of these loci 
(RPP1, RPP2, RPP4 and RPP7) were mapped (Tor et al., 1993), along with a further 
locus RPP5 (Parker et al., 1993). Three loci were apparently clustered together on 
chromosome four. There was also evidence for the existence of different alleles at a 
single locus, although corresponding crosses between pathogen isolates differing in 
specificity are required to confirm whether different alleles at closely linked loci might 
explain such results. A genetic model postulating the existence of complementary 



109 

Table 23. Inheritance of resistance in cauliflower to P. parasitica using 
Palermo green model (Reprinted from N.A. Moss, I.R. Crute, J.A. 
Lucas and P.L. Gordon. 1988. Requirements for analysis of host- 
species specificity in Peronospora parasitica (downy mildew). 
Cruciferae NewsLetter 13:114-116, by permission of the author and 
the publisher) 



P. parasitica 


Virulence 


Resistance phenotype 


Approx 


. % resistant 


Isolate 


r 




- 


Rl - 


Rl 


seedlings observed 




Al 


A2 


- 


R2 


R2 






P005 


1 


2 


+ 








95 


P015 


1 


- 


+ 


+ 


- 




80 


P018 


- 


2 


+ 


+ 


- 




80 


P006 


- 


- 


+ 


+ + 


+ 







% Phenotype in 
















seed stock 






5 


15 15 


65 







A = avirulence gene, R = Resistance gene, + = susceptibility, - = resistance 



recognition loci in the fungus designated ATR- A. thaliana recognition has been 
proposed (Holub et al., 1994). These results provide support for the gene-for-gene model 
of specificity in crucifers downy mildew. The interaction phenotypes between different 
host lines and pathogen isolates reveal a degree of complexity in the system, with 
partial dominance, epistasis and gene dosage effects. 

e. Biochemical basis of resistance 



The presence and absence of natural biochemical compounds like glucosinolates 
and other phenolic compounds play a significant role in providing resistance to the host 
plant. There is a correlation between high levels of flavour volatiles (e.g. 
allylisothiocyanate) released by tissue damage, and the limitation of fungal growth in 
both wild and cultivated Brassica lines. In cultivated brassicas, breeding has resulted 
in reduced levels of flavour volatiles with a consequent reduction in their general 
resistance to P. parasitica. The resistance to P. parasitica in the cabbage cultivar 
January King may be attributed to the high concentration of allyl- 
isothiocyanate(Greenhalgh and Mitchell, 1976). 



110 



Table 24. Response of groups A, B, C, D and E of Brassicajuncea accessions 
and of one accession of B. napus at the cotyledon stage to infection 
with four isolates of Peronospora parasitica (Nashaat and Awasthi, 
1995) 



Host 
category* 



Disease Index 



Isolates of P. parasitica 
IP01/IP02 P003 Rl 



B. juncea 



A (4) 

B(2) 

C(19) 

D(5) 

E(l) 



1 

2-3 

6-8 

6-8 

6-7 



1 

1 

1 

2-3 

5-6 



B. napus 



Ariana 



7-8 



7-8 



c ) Number of accessions of each group in parenthesis. 



The incidence and severity of downy mildew is positively correlated with 
glucosinolate concentration in seeds of oilseed rape (Rawlinson et al., 1989). In oilseed 
rape, downy mildew severity is lower on cultivars with high concentration of 
glucosinolate (>100 /^mol g" 1 ) and greater on those with a lower concentrations (<15 
^mol g" 1 ). In all cultivars grown in the UK incidence is lower in mid-February when 
glucosinolate products in the leaves reaches a maximum level (Anonymous, 1985). 

A large number of B. napus species with different glucosinolate and erucic acid 
contents have been screened for resistance to four isolates of P. parasitica at the 
cotyledon stage (Nashaat and Rawlinson, 1994). Two groups of accessions with 
different resistance factors were identified. The first group was different from that of 
the cultivar Cresar which has an isolate specific gene for resistance to P. parasitica, 
and the second group was identical to that of Cresar. There was moderate to full 
susceptibility at the cotyledon stage but no clear differential response to any of the 
isolates. Those with high glucosinolate and high erucic acid content were significantly 
less susceptible than those with high glucosinolate and low erucic acid, or, low 
glucosinolate and low erucic acid content. 



In preliminary experiments on the effect of treatment with abiotic elicitors on 
disease reaction in oilseed rape seedlings, salicylic acid and methyl jasmonate reduced 



Ill 



Table 25. Examples of a successful selection for putative homozygous 
resistance response to Peronospora parasitica from a 
heterogeneous starting population of Brassica juncea at the 
cotyledon stage (Nashaat and Awasthi, 1995) 



B. juncea' 



IP01 



Isolates of P. parasitica 
Disease Index (Standard deviation, n = 24) 

IP02 P003 Rl 



Kranti 


3.0 (3.0) 


4.1 (3.6) 


1.0(0.0) 


1.0 (0.0) 


RES-BJ01 


1.0 (0.0) 


1.0 (0.0) 


1.0 (0.0) 


1.0 (0.0) 


Krishna 


5.4(2.5) 


5.7 (2.5) 


1.0 (0.0) 


1.0 (0.0) 


RES-BJ02 


1.0 (0.0) 


1.0 (0.0) 


1.0 (0.0) 


1.0 (0.0) 


Varuna 


6.1 (1.8) 


7.1 (2.2) 


1.0 (0.0) 


1.0(0.0) 


RES-BJ03 


1.0 (0.0) 


1.0 (0.0) 


1.0 (0.0) 


1.0(0.0) 



*) RES-BJ01, RES-BJ02 and RES-BJ03, lines selected from seedling populations of 
Kranti, Krishna and Varuna, respectively. 



the severity of infection when cotyledons were subsequently inoculated with P. 
parasitica (Doughty et al., 1995). 

f. Lignification of host cells 

Lignin formation was observed in cell walls of parenchyma of Japanese radish 
root infected with downy mildew fungus. The observed lignin was mainly composed 
of guaincylpropane units and apparently differed from syringyl lignin which was 
present in vessels of the healthy tissues. Different pathways for lignin biosynthesis 
in the healthy and the diseased tissues were proposed (Tables 26, 27, Fig. 59-61) by 
Asada and Matsumoto (1972). Specific isoperoxidases synthesized de novo in diseased 
tissues were presumed to play an important role in the formation of the guaincyl lignin 
(Ohguchi et al., 1974; Ohguchi and Asada, 1975). Lignin was also formed in cell walls 
of parenchyma of Japanese radish root which was infiltrated with 700 x g supernatant 
of homogenate of downy mildew infected tissues. It began to form about 12h after 
infiltration of the homogenate (Matsumoto et al., 1978). The effective component in the 
homogenate for the induction of lignification is dialyzable, which is highly water 
soluble and seems to resemble monilicolin A (Asada et al., 1975). The lignification 



112 




260 280 

WAVELENGTH (nm) 



Fig. 59. The UV absorption spectra of the diseased parenchyma cell wall (A), the vessel wall (B), and the healthy parenchyma cell 
wall (C) of the Japanese radish root (Reprinted from Y. Asada and I. Matsumoto. 1972. The nature of lignin obtained from 
downy mildew-infected Japanese radish root. Phytopathol. Z. 73: 208-214, by permission of the authors and the publisher 
Blackwell Wissenschafts-Verlag GmbH, Berlin, Germany). 




WAVELENGTH (nm) 



Fig. 60. UV absorption spectra of the authentic compounds (A) and the degradation products (B) obtained from the extraction of paper 
chromatograms. I = P - hydroxybenzaldehyde; II = vanillin; III = syringaldehyde (Reprinted from Y. Asada and I. 
Matsumoto. 1972. The nature of lignin obtained from downy mildew-infected Japanese radish root. Phytopathol. Z. 73: 
208-214, by permission of the authors and the publisher Blackwell Wissenschafts-Verlag GmbH, Berlin, Germany). 



113 



tran s-p-Coumarx tc, Cinnamoyl esl«*t 
carbnxyl activation 
hydroxylase 

X "^<o> 



»o(o> 



CHCHCOOE 



Px 



CHCH OH "* 



^ p-OH-ph*nyl aoiety— 1 



p-Coumaf.y I qitft 

— phenol a se 

— o-methy 1 trimfiriie 



p-Cou«aryl alcrfl.-t 



HOf O) CHCHCOOE 



Py 

» H 3 C0 - 



"°<§) 



CHCHCH OH 



Feruloyl osl« Coniferyl alcnh.l 

■ hydroxylase 
n-avthyl transferase 



HjCO >l"^ 

IK>/0\cHCHCOOE 
H CO 

Sinapoyl ntii 



H„CO 



3 „~<§> 



J 



> CHCHCH OH 
H 3 CO~ 

Sialyl \lcohol 



Guaiacyl nioicty ^3: 



Syrinrjyl aoiety 



1 



^•Dis<v\scd lignin 



J 



Healthy li<snla 



Fig. 61. Suggested pathway of lignin biosynthesis in healthy (full lines) and diseased (broken lines) plants. Px, Py, Pz: Peroxidase 
isoenzymes x,y,z (Reprinted from Y. Asada and I. Matsumoto. 1972. The nature of lignin obtained from downy mildew- 
infected Japanese radish root. Phytopathol. Z. 73: 208-214, by permission of the authors and the publisher Blackwell 
Wissenschafts-Verlag GmbH, Berlin, Germany). 



114 

inducing factor (LIF) plays a significant role in the induction of systemic resistance. 
Resistance to P. parasitica was induced in roots of susceptible radish cultivars when 
they were preliminarily inoculated with the pathogens or wounded. An increase in L- 
phenylalanine ammonia lyase (PAL) activity and lignification of cell walls occurred in 
these tissues after challenge inoculation with downy mildew. Histochemical 
observations indicated that the cell walls were lignified in the tissues beyond the site 
of fungal attack (Matsumoto and Asada, 1984). High peroxidase activity was located 
around the lignified cell walls (Asada and Matsumoto, 1969; Ohguchi et al., 1974). The 
higher amount of lignin accumulation was present in the middle portion of the cell wall 
(Asada and Matsumoto, 1971). Following infection of radish by P. parasitica the 
deposition of lignin in host cell walls may have a role to play in non-specific limitations 
of the growth of biotrophic fungi. Ohguchi and Asada (1975) have defined the pathways 
and enzymes involved in lignin biosynthesis in radish following infection by P. 
parasitica. 

g. Sources of resistance 

Differential host resistance to isolates of P. parasitica has been identified in B. 
campestris, B.juncea, B. napus, B. oleracea and R. sativas (Bonnet and Blancard, 1987; 
Lucas et al., 1988; Nashaat and Rawlinson, 1994; Nashaat and Awasthi, 1995; Nashaat 
et al., 1997; Silue et al., 1996). Sources of resistance to Peronospora parasitica in 
different host species of crucifers identified from different countries of the world are 
given in Table 28. 



10. BREEDING FOR DISEASE RESISTANCE 

Plant breeding offers one method for controlling diseases and has obvious 
advantages if successful. As with other traits, the breeder's task is firstly to find 
sources of disease resistance and effective ways to screen for resistant genotypes. The 
trait has then to be transferred into a useful cultivar or hybrid (Buzza, 1995). Transfer 
of resistance among crucifers and from other species is possible by using conventional 
and biotechnological techniques: 

a. Germplasm evaluation for sources of resistance at national and international 
levels. 

b. Selection for disease resistance through (i) pure line selection, (ii) mass selection, 
(iii) modified recurrent mass selection, and iv) recurrent selection. 

c. Breeding for disease resistance by increasing the level of resistance through i) 
multiple crosses, ii) recurrent selection, iii) dialle crossing, and d) selective mating 
system 

d. Transfer of resistance by i) intraspecific pedigree, backcross and modified 



115 



Table 26. Amounts of degradation products by alkaline nitrobenzene 

oxidation of the isolated lignin (Reprinted from Y. Asada and I. 
Matsumoto. 1972. The nature of lignin obtained from downy 
mildew-infected Japanese radish root. Phytopathol. Z. 73:208- 
214, by permission of the authors and the publisher Black we 11 
Wissenschafts-Verlag GmbH, Berlin, Germany) 



Lignin 



Product 



Amount (%) 



Ratio 





p-Hydroxybenzaldehyde (H) 


0.39 


V/H 6.05 




Vanillin (V) 


2.36 


S/H 3.90 


Healthy 


Syringaldehyde (S) 


1.52 


S/V 0.64 




Total 


4.27 






p-Hydroxybenzaldehyde (H) 


0.99 


V/H 2.75 




Vanillin (V) 


2.72 


S/H 0.0 


Diseased 


Syringaldehyde (S) 


0.00 


S/V 0.0 




Total 


3.71 





recurrent mass selection methods, and ii) interspecific genome substitutions, 
chromosome substitutions and gene introgression. 

e. Transfer of resistance through mutation breeding. 

f. Use of biotechnological and genetic engineering techniques such as i) genome 
manipulation, ii) manipulation of cytoplasmic genomes, iii) use of transformation 
and foreign gene expression techniques, and iv) embryo rescue techniques for wide 
hybridization. 



11. DEVELOPMENT OF RESISTANCE TO FUNGICIDES 



The existence of metalaxyl resistant strains of P. parasitica has been reported 
(Crute, 1984; Crute and Gordon, 1986; Brophy and Laing, 1992). Metalaxyl resistant 
isolates were also shown to be cross-resistant to furalaxyl and ofurace, two related 
phenylamide fungicides. A differential degree of insensitivity to two related 
phenylamide fungicides has been demonstrated (Table 29) (Moss et al., 1988). 
Metalaxyl was more active against a sensitive isolate (P005) than cyprofuran but the 



116 

Table 27. Elemental compositions and empirical formulae of the isolated 
lignins and the related compounds (Reprinted from Y. Asada 
and I. Matsumoto. 1972. The nature of lignin obtained from downy 
mildew-infected Japanese radish root. Phytopathol. Z. 73:208-214, 
by permission of the authors and the publisher Blackwell 
Wissenschafts-Verlag GmbH, Berlin, Germany) 



Lignin 


C(%) 


H(%) 


OCH 3 (%) 


Formula 


Healthy root 


64.38 


7.27 


18.93 


^9"l0.3^'2.20^^- / "3n.l6 


Diseased root 


63.47 


5.74 


12.58 


^- / 9-"-8.33^2.80^^- / -"-3^0.75 


Birch*) 


58.82 


6.49 


21.51 


CgHg 0302.77(00x13)! 58 


Spruce*) 


63.48 


6.35 


14.84 


^9-"-8.83^'2.37^OL/rl3) 96 


DHP**) 


64.00 


6.00 


16.90 


^9"8.2l02.5o(OCH 3 ) 1 2 


Coniferyl ale. 


66.67 


6.67 


17.22 


C^O^OCHg)! 



*) From BJORKMAN and PERSON (1957). 

**) Dehydrogenation polymerization product, from NOZU (1967). 

The formulae were obtained from the following equations. 

GjH.OytOCHg), where 



x = 



1.008 X 3 
108.09 TH (%) - 31.035 



X OCH ? (%)} 



1.008D 



z= 108.9 X OCrL (%) 
31.035D 



16 



y= 108.09 TO (%) - 31.035 X OCH 3 (%)1 

16D 



12.01 
D = C (%) -- 31.035 x OCH, (%) 



converse was true with an insensitive isolate (P006). The inheritance of fungicide 
insensitivity to P. parasitica may reveal the true picture of genes controlling this 
phenomenon. 

12. LABORATORY AND FIELD TECHNIQUES AND BIOASSAYS 



a. Culturing of Peronospora parasitica 



Table 28. Sources of resistance to Peronospora parasitica 



117 



Host species/genotypes 



References 



Brassica alba (white mustard) 
All Indian accessions 

B. carinata (Ethiopian mustard) 

All Indian accessions 

HC1 

B. campestris 

candle 

B. campestris var toria (Toria) 
IB - 586 

B. campestris var yellow sarson (Yellow Sarson) 
YST-6 

B. campestris var brown sarson (Brown Sarson) 
BS-15 

B.juncea (Indian mustard) 

PI 340207, PI 340218, PI 347618 

Domo, RC 781, EC 126743, Zem, YRT 3, 45, 72 

PR 8805, RN 248, EC 129126-1, PC 3 

RESBJ-01, RESBJ-02, RESBJ-03 

B. oleracea var. botrytis (cauliflower) 

Igloo, Snowball y, Dok Elgon, RS-355 

PI 181860, PI 188562, PI 189028 (MR), PI 204765, PI 204768, 

PI 204772, PI 204773, PI 204775, PI 204779, PI 241612, 

PI 264656, PI 291567, PI 373906, PI 462225 (MR) 

KPS-1 

PI 231210, PI 189028 

B. oleracea var capitata (cabbage) 

January King 

Balkan 

Spitz Kool 

PI 246063, PI 246077, PI 245013 

Tromchuda cabbage "Algarvia" (ISA 207) 

PI 245015, Geneva 145-1 

B. oleraceae var accephala gr. ornamentalis 
(Decorative cabbage) 



Saharan, 1992a, b 



Saharan, 1992a, b 
Saharan, 1996 

Saharan, 1992a, b 



Kolte & Tewari, 1980 



Kolte & Tewari, 1980 



Kolte & Tewari, 1980 



Ebrahimi et al., 1976 
Saharan, 1992a, b 
Saharan, 1996 
Nashaat and Awasthi, 1995 

Kontaxis et al., 1979 



Thomas and Jourdain, 1990 
Sharma et al., 1991 
Hoser-Krause et al., 1991 



Greenhalgh & Mitchell, 1976 

Elenkov, 1979 

Verma & Thakur, 1989 

Hoser-Krause et al., 1991 

Caravalho & Monteiro, 

1996 

Sherf and Macnab, 1986 

Vitanova, 1996 



B. oleracea (Broccoli) 



118 



calabrese, Grand Central 

PI 231210, Italian Green Sprouting 

Hyb. 1230 (Moran), Green surf (Moran), 

2804 (Qualisal), Hyb. 2805 (Qualisal), 

Hyb. 2803 (Qualisal), GSV 82-4310 (Goldsmith), 

XPH 1117 (Asgrow), Hyb. 288 (Moran), 

AVX 7631 (Sun Seeds). 

PI 263056, PI 263057, PI 3573, PI 3574. 

PI 418984, PI 418985, PI 418986, PI 418987 

PI 418988 

OSU CR 2 to OSU CR 8 

Citation, Excalibur, Nancy 



Natti et al., 1956 

Natti, 1958 

Laemmlen & Mayberry, 1984 



Hoser-Krause et al., 1991 



Baggett and Kean, 1985 
Sherf and Macnab, 1986 



B. napus (Rape) 

Hg Vestal 

Eurora, Janetzki, Kubla, Lesira, 

Mogul, Primar, Rapot, Rapara, Sinus 

cultivar 78-22 

Cresor 

PI 199949, PI 263056 

Gulivar, Midas, Tower 

RES 01-1-4, RES-02, RES-26 

HNS3, HNS4, GSL 1, GSL 1501 



Jonsson, 1966 
Dixon, 1975 

Chang, 1981 

Kluczewski & Lucas, 1983 
Thomas & Jourdain, 1992 
Saharan, 1992a, b 
Nashaat et al., 1995; 1996 
Saharan, 1996 



B. chinensis (Chinese cabbage) 

Bau chin 26, PHW 64707, PHW 64710, 

PHW 64722, PHW 64620 

Hyb. 77M(3)-27, Hyb. 77M(3) - 35 

Hyb. 82-46, Hyb. 82-46R, Hyb. 82-156, 

Hyb. 82-157 



Niu et al., 1983 
Anonymous, 1987a, b 



B. nigra (Black mustard) 
PI 199948 



Thomas & Jourdain, 1992 



B. napa 

PI 418984, PI 418988, PI 418987, PI 418988 



Thomas & Jourdain, 1992 



B. rapa subsp. rapifera 

Long Blanc de croissy, Stanis, Jaune Boule d'or 

Raphanus sativus (Radish) 

Okura 

Tokinoshi (All season) 

Bamba, Noir Lon d'Horloge, Rave a Forcer 



Silue et al., 1996 

Shiraishi et al., 1974 
Bonnet & Blancard, 1987 
Silue et al., 1996 



Cheiranthus cheirii (Wallflower) 
Convent Garden blood Red 



Greenhalgh & Dickinson, 1975 



119 

Table 29. Responses of phenylamide sensitive and insensitive isolates of P. 
parasitica to phenylamide fungicides (Reprinted from N.A. Moss, 
I.R. Crute, J.A. Lucas and P.L. Gordon. 1988. Requirements for 
analysis of host-species specificity in Peronospora parasitica 
(downy mildew). Cruciferae NewsLetter 13:114-116, by permission 
of the authors and the publisher) 



Compound Isolates Fungicide 

0.05 0.05 



yug/ml Factor of Insensitivity 
5.0 50.00 



Metalaxyl 


P005 
P006 


32 a 
94 



89 



100 



83 



xlOOO 


Cyprofuran 


P005 
P006 


84 
100 


79 

94 



78 



3 


xlO 



a. Figures are reciprocals of mean latent periods (time from inoculation to sporulation 
expressed as a percentage of the untreated control). 



The biotrophic nature of this pathogen implies a sophistication of nutritional 
requirement which can not be met upon the death of the host plant. P. parasticia was 
cultured on disinfected slices of swede root where aerial growth of mycelium was 
observed with conidiophores, conidia, antheridia, oogonia and oospores (Guttenberg 
and Schmoller, 1958). Ingram (1969) successfully established and maintained cultures 
of the fungus on callus tissues derived from (a) a mature leaf of cabbage, (b) a mature 
root/hypocotyl of rape, and (c) a seedling hypocotyl of swede. The infected calluses were 
incubated either at 22 °C in the dark, or at 15 °C with 12h fluorescent illumination 
photoperiod. To maintain the dual culture of the callus and the pathogen, sub- 
culturing through transfer of an explant of infected callus to fresh uninfected callus 
was necessary every 14 - 21 days. Calluses derived from the root/hypocotyl of rape 
grew faster than those from cabbage leaf, and were transferred directly when 
subculturing. About four weeks after inoculation of the rape callus, small nodules of 
new tissue were developed on the infected callus. These nodules or the whole callus 
were successfully maintained when transferred to fresh culture medium. Conidia of 
P. parasitica were produced on infected callus tissue maintained at 15 °C and 12 
h/photoperiod but production was much lower at 22 °C in the dark. Such conidia were 
used to infect detached cotyledons or leaf callus of cabbage or rape. 

Attempts were made with limited success by Guttenberg and Schmoller (1958) to 
culture the fungus in the absence of living plant tissue. They obtained visible mycelial 



120 

growth in filter, sterilized swede juice, but it ceased after 3 days when a yellow 
discoloration appeared suggesting that the medium was chemically unstable. Very 
limited mycelium was developed on swede seed-glucose agar and maize decoctions- 
glucose agar, but more success was achieved with an agar medium containing 2% beer 
wort + 0.1% phosphate in which hyphae and conidiophores were developed within and 
outside the agar substrate. Similar, but less growth was achieved in oatmeal agar and 
rice starch agar. 

Asada and Ohguchi (1981) studied the behavior of downy mildew fungus of 
Japanese radish on modified Knop's medium. An isolate of P. parasitica from naturally 
infected leaves was cultured on modified Knop's medium and 0.1% streptomycin, using 
infected slices of radish root tissue as inoculum. Vigorous hyphal growth was observed 
spreading into the medium and numerous haustorium like bodies were formed. The 
production of which was favoured by low agar concentration, low pH(4) and high 
sucrose concentration (50 g/1). Growth was ceased 2 weeks after placing the tissue 
slices on the medium. 

The response of P. parasitica to a liver medium was studied by McMeekin (1981). 
Washed, autoclaved 2mm cubes of liver were placed in a 9 cm petri dish and covered 
with 20 ml of a mixture of 0.01% tryptone (Difco: pancreatic digest of casin), 0.04% 
K2HP0 4 and 2% agar: a 1/10 dilution of the K2HP0 4 and tryptone. Ten ,ug/ml strepto- 
mycin (Calbiochem) were added to this medium to control bacterial growth. When the 
agar was solidified, drops of a suspension of conidia in water were placed on the piece 
of fiver. The plates were incubated at 18°C. After 4 days the germ tubes in the control 
(without liver) disintegrate, but in a 5 mm diam zone around the liver pieces, 90% of 
the germ tubes grow towards the liver. They grow from the surface towards the bottom 
of the medium and formed large swellings or lobes within the agar. The lobed germ 
tubes reached their maximum size after 4 days. When the plate was flooded with 
sterile distilled water, either 4 or 7 days after the beginning of conidial germination, 
the lobes on the germ tubes maintained the same size. 

b. Maintenance of P. parasitica isolates 

Isolates of P. parasitica were maintained separately on cotyledons obtained from 
6-days old seedlings, raised in soilless compost in a modified plant propagtor (35.5 cm 
x 21.6 cm x 18 cm), sited in the glasshouse (Nashaat and Rawlinson, 1994). The 
propagator was supplied with continuous filtered (spore-free) moist air at 18+2°C 
through a central flue conducting air from beneath the propagator to exhaust at two 
adjustable ventilators on the cover and the junction between the cover and the base 
(Jenkyn et al., 1973). Supplementary light was given to maintain a 16h photoperiod. 
Cotyledons and a short length of hypocotyl were detached and transferred to folded 
filter paper (Whatman 12.5cm, 113v) supports in glass jars (8cm diameter, 7 cm depth) 



121 

containing 20 ml sterile distilled water. Cotyledons were then inoculated in a sterile 
air flow with 5/A of conidial suspension on each half cotyledon with the aid of a 
micropipette. Conidial suspensions were prepared by tapping infected cotyledons to 
dislodge conidia into sterile distilled water; this minimized bacterial contamination. 
After inoculating the cotyledons, the glass jars were covered with clear plastic lids, 
sealed with parafilm and incubated in a growth cabinet at 16°C under 70/^E/m s 2 " 
irradiance with a 16h photoperiod for 7 days after which peak sporulation occurred. 

c. Germplasm screening and evaluation 

Genotypes are grown in propagators, (as described under isolates maintenance in 
subsection 12b) except that two adjacent 5 cm 'Jiffy-pots' for each line or cultivar are 
used as pots. The pots are placed on capillary matting to ensure a uniform water 
supply. Each propagator contains up to 13 accessions arranged as two randomized 
blocks (propagators) with each accession occurring only once in each propagator. 
Initially, nine to 15 seedlings per accession are grown in each propagator and these are 
thinned 6-days after sowing to 6 to 10 to decrease variability in growth. Sowing dates 
are staggered to produce seedlings at the required growth stage for inoculation at the 
same time. The average times required under these conditions to reach fully expanded 
cotyledons, first, and second true leaves are 7, 16, and 22 days, respectively (Nashaat 
and Rawlinson, 1994) . 

Seedlings are inoculated by spraying them to run-off with a suspension of conidia 
(2.5 x 10 conidia/ml). The propagators are sealed after inoculation to allow the relative 
humidity to rise to 100%, and then incubated in growth cabinets under the conditions 
described for isolates maintenance. Infection phenotypes are recorded 7 to 9 days after 
inoculation (on cotyledons and leaves, respectively) using a 0-9 scale (Nashaat and 
Rawlinson, 1994). 

According to Williams and Leung (1981), single seeds are grown in 12-pack pots 
and when the cotyledons have expanded, after five to seven days, single 0.01 to 0.02 
ml drops of a freshly collected conidial suspension containing approximately 10 5 
conidia/ml are placed on each half of the two cotyledons using a finely tipped glass 
pipette. As plants of each 12-pack are inoculated they are placed in glass or plastic 
boxes containing a 1 to 2 cm depth of warm water in the bottom. A tight fitting cover 
is placed on the box after the box is filled with plants. The box is then placed in a 
darkened incubator at 20°C for 8 to 16h. The atmosphere in the box will maintain the 
droplets on the cotyledons during which time germination and penetration will occur. 
After incubation, plants are placed on a lighted greenhouse bench at 20 to 25°C for five 
days then returned to humidity boxes at 20°C for 16-24h. Upon removal from the 
humid atmosphere, susceptible plants will have a profuse growth of P. parasitica 
conidiophores on the abaxial sides of the cotyledons, whereas resistant plants will 



122 

exhibit varying degrees of sporulations and tissue necrosis which can be evaluated on 
a 0-9 scale (Fig. 62). A rating of 1 is given to rapidly occurring (24 to 48h) 
hypersensitive necrotic flecking, without visible sporulation, found immediately under 
the droplets. Interaction phenotypes representing host-pathogen compatibility are 
expressed as increasing degree of sporulation on the abaxial side of the cotyledons and 
decreasing degree of necrosis associated with tissue colonization. It is important to use 
freshly produced inoculum collected by washing off spores from leaves with distilled 
water. Older plants may be inoculated by atomizing a suspension of conidia on the 
foliage and holding them at 100% RH for 8-16h. 

Knight and Furber (1980) used descriptive key for the assessment of downy 
mildew disease prevalence and severity on winter oilseed rape varieties. A group of 
plants (10 plant/sample) chosen at random from a number of observation site assessed 
were as follows: 



Percentage disease 



Host - pathogen interaction 



0.0 
0.1 

1 
5 



10 
25 

50 

75 
100 



No infection 

Traces of infection generally confined to lower leaves. 

One plant in ten or fewer with lesions 

Some plants infected, but one or two lesions per plant 

Most plants affected with about 5% of the lower leaf 

area affected 

Most plants with about 10% of the lower leaf area 

affected. Up to 5% infection on the upper stem leaves 

and bracts 

About 25% of the lower leaf area affected. Leaf area 

may appear to be half affected and half unaffected. 

Infections frequent on upper stem leaves and bracts 

(up to 10% area affected) 

About 50% of the leaf area affected. Affected area 

appears to be greater than unaffected 

About 75% of leaf area affected. Very little unaffected 

tissue observable 

100% of leaf area affected 



123 



ADAXIAL 



AB AXIAL 




HYPERSENSITIVE 
NECROSIS 



— ~_ TISSUE 

"2T" wtrosis 



Jy Y = SPORULATJON 

Y yY 



Fig. 62. Rating scale for downy mildew (Peronospora parasitica) interaction phenotypes on Chinese cabbage (Reprinted 
from P.H. Williams and H. Leung. 1981. Methods for breeding for multiple disease resistant Chinese cabbage. IN: 
Chinese cabbage. Proc. 1st Intern. Symposium, N.S. Talekar and T.D. Griggs (Editors): pp. 393-403, by permission 
of the authors and the publisher the Asian Vegetable Research and Development Center, Shanhua, Taiwan). 



124 

d. Preservation of P. parasitica 

Brassica leaves infected with the pathogen were collected and conidia from such 
infected material were inoculated on cotyledons of a susceptible Brassica variety (Paul 
and Klodt-Bussmann, 1993). Cotyledons are put into plastic boxes on moist filter 
paper and incubated at 15°C and 70-80% RH. Under these environmental conditions 
conidia and conidiophores of the pathogens are ready to be harvested 6-days after 
inoculation. For each isolate 5 cotyledons colonized with fresh conidia were collected 
in a glass vial in 10% (v/v) glycerine which serves as a cryoprotectant in the suspension 
medium. For each isolate, 6 glass vials are filled with the conidial suspension and 
immediately stored in a freezer at -21°C. After a storage interval, samples are taken 
out from the freezer and thawed at room temperature (20°C). After thawing for 10 
minutes, 2 ml of the conidial suspension are added to each petri-dish which contains 
15% (w/v) water-agar and the percentage of conidial germination is assessed after 24h 
at 15°C. After a storage period of 8-days it was found that the highest germination rate 
of 73% occurred using 10% (v/v) glycerine. 

To culture and preserve Japanese Radish downy mildew fungus, slices (1cm) of 
Japanese radish root cv Miyashige, were inoculated with conidia of the fungus collected 
from naturally infected leaves and incubated at 20°C for 6-8days (Ohguchi and Asada, 
1981). Conidia produced on the infected slices were then collected to make a 
suspension and were used to inoculate other healthy slices. Numerous oospores are 
observed in these sliced tissues 6 days after inoculation. 

The conidial viability of P. parasitica derived from cabbage was longest, up to 130 
days, when the spores were stored in air dried soil at a constant temperature of 5°C. 
Conidia kept at -25°C and relatively dry on leaf disks (air dried at 20°C) maintained 
a relatively high rate of germination after 1 year or longer (Krober, 1970, 1981). 

The pathogen is usually preserved by storing few sporulating cotyledons in small 
vials with tight lids inside a deep freezer (-25 to -30°C) for up to 6 months. 
Temperature fluctuations should be avoided during this period. For reviving the 
pathogen, the vials containing the sporulating cotyledons are taken out of the freezer 
and, with their lids kept tightly on, transferred immediately to a container containing 
icy water. The temperature of the container and its content is then allowed to rise 
gradually, within l-2h, to 15°C. Thereafter, the vials are taken out of the container 
and conidial suspension for inoculating fresh cotyledons is prepared in the normal way 
(Nashaat and Rawlinson, 1994). 

e. Artificial inoculation of excised cotyledons 

Cotyledons of radish, 9 days after sowing, were placed face downwards on a damp 



125 

filter-paper in a transparent plastic box (Bonnet and Blancard, 1987). The box was 
then put in a growth chamber at 20°C day and 18°C night, relative humidity of 90%, 
and illumination of 2000 lux for 12h. With a micropipette, 50 [A of P. parastica conidial 
suspension (16,000 sp/ml) were placed on the abaxial surface of the cotyledons. After 
5 days, the conidia were collected by rubbing the cotyledons with a brush into 2 ml of 
water; the concentration of the suspension was then measured and adjusted using a 
hemacytometer. When 15-day-old plants were inoculated, an excellent correlation was 
observed between the number of conidia and symptoms on leaves. 

f. Propagation of P. parasitica on cotyledons or true leaves of Japanese 
radish seedlings 

Cotyledons and the true leaves of radish seedlings were subjected to hot water 
(50°C) treatment, or the roots were cut off to weaken resistance to P. parasitica 
(Ohguchi et al., 1989). Each of the 7-11 days-old cotyledons of cvs. Awa-ichigo, 
Sarakamuri and Daimaru-Shogoin, which had been treated with hot water, were put 
in a test tube (2.8 x 19 cm) containing 15 ml of distilled water. The upper surfaces of 
these cotyledons were inoculated with drops of conidial suspension of the fungus. The 
percentage of conidiophore formation on the cotyledons grown for one week at 20°C, 
1000 lux after inoculation was highest on the 11 day old cotyledons from cv. Diamaru- 
Shogoin treated with hot water for 60 seconds. On the true leaf of 3 week old 
seedlings, the percentage was highest in cv Sarakamuri treated with hot water for 30 
seconds. Since the cotyledons of cv. Shirokubi-miyashige, Heian-tokinashi and 
Daimaru-Shogoin, which had been grown for 4 to 6 days in a growth chamber (25°C; 
5,000 lux) were very susceptible to hot water treatment, their roots were cut off. Cut 
surfaces of hypocotyls were wrapped with cotton wetted with sterilized distilled water 
or a modified Knop solution in order to keep the cotyledons from withering. The lower 
surfaces of cotyledons were inoculated with the suspension of conidia. The percentage 
of the conidiophore formation was highest on the 6-day-old cotyledon of cv. Shirokubi- 
miyashige. In the case of cv. Daimaru-Shogoin, the 4-day-old cotyledon was best 
suited. A dark treatment of 18 h of the infected cotyledons on the 6th day after 
inoculation stimulated conidiophore formation following synchronized formation of the 
conidia. Also, many conidiophores were formed on the infected cotyledon when moved 
into an incubator at 20°C after being stored in a refrigerator at 5°C for two weeks after 
the 3rd day after inoculation (Ohguchi et al., 1989). 

g. Laboratory tests of fungicides 

A susceptible cultivar of Brassica species must be used for the maintenance of P. 
parasitica (Channon and Hampson, 1968). Sow seeds of susceptible cultivar in boxes. 
Detach cotyledons bearing 4-5 mm petiole from the seedlings and lay them in a single 
layer on sterilized moist, crinkled filter paper in transparent plastic boxes. Add 



126 

sufficient sterilized tap water to maintain the filter papers adequately moist. Ten to 
fourteen day-old cotyledons are suitable for maintaining the cultures. Obtain conidia 
from an actively sporulating fungus on leaf or cotyledon. Seedlings can be inoculated 
with the aid of a small paint brush or spraying, or by dipping the cotyledons in the 
spore suspension. After inoculation, incubate the boxes of cotyledons in growth cabinet 
at 15°C with illumination. Supplementary light (400w mercury fluorescent lamps, 3 
3/4 ft above the boxes and each illuminating an area of just under 11 sq ft) for 12h per 
day is essential for the survival of both infected and uninfected cotyledons. 

To test the protectant action of fungicides, 10-14 day old seedlings are cut off at 
soil level and placed in Weldmesh Racks (14 seedlings per rack) with cut ends of the 
stems immersed in water in an enamel dish which support the racks (Channon and 
Hampson, 1968). Atomize 2 ml of the test chemical on the upper surfaces of leaves of 
seedlings in the racks. On the following day cut off the leaves with petioles and place 
on moist filter paper in three to four plastic boxes (3 1/3" x 1 7/8" x 7/8"). Inoculate 
these leaves with a drop (0.01 ml) of spore suspension containing approximately 1000 
conidia and incubate at 15°C in an illuminated incubator. Record the number of leaves 
showing sporulation. 

h. Fungicide resistance assay 

An assay for resistance and sensitivity of P. parasitica to metalaxyl can be made 
using cauliflower seedlings of cv. Lawyna (Crute et al., 1985). Nutrient solution (25 
ml) amended metalaxyl (Ridomil 25 W.P.) at a range of concentration of up to 100 
Mgml' 1 was contained in 7 cm diameter glass crystallizing dishes and absorbed in an 
equal volume of vermiculite. Seed was sown into the dishes (30 - 40 per dish) and 
placed in a temperature controlled growth room (15°C, 12h photoperiod, 100 //Em^S" 1 ). 
To avoid problems with vapour activity, each dish was contained within a plastic 
"treacle pot'. Seedlings at the cotyledon stage, 7-10 days after sowing, were inoculated 
with the conidial suspension of the fungus and incubated under the same conditions. 
Observations were recorded for the presence or absence of sporulation 5-10 days after 
inoculation. A standard metalaxyl sensitivity isolate which was completely inhibited 
at O.Ol/xgml' 1 was included in all tests. A modification of the method using cauliflower 
seed treated with metalaxyl (Ridomil 25 WP) at a rate of 1 g a.i. per kg clearly 
discriminated between resistant and sensitive isolates. The bioassay of the plant 
material revealed 15-20 fj.g 'metalaxyl equivalents' per g fresh weight in seedlings 
during the course of the test. Resistant isolates sporulated profusely on seedlings 
grown from treated and untreated seed while sensitive isolates only sporulated on the 
later (Crute et al., 1985). 



127 

i. Measuring systemic infection by the downy mildew pathogen 

According to McMeekin (1971), seeds of brassicas were first surface sterilized in 
sodium hypochlorite (10% Commercial Clorox) for 5 minutes, and then placed about 1.5 
cm apart on either 1-2% agar or sterilized glass wool moistened with distilled water in 
the bottom of a moist chamber (McMeekin, 1971). They were germinated in the dark 
at 20°C. Seven days later most seedlings were about 2 cm tall. The cotyledons and 
roots were excised from these seedlings. The hypocotyl, relatively free of starch 
granules and chloroplast, was placed with one end in 10 ml of test solution at the 
bottom of a Petri dish (5 cm diameter). Ten or more hypocotyls were kept upright in 
the dish by pushing them through a double layer of cheesecloth stretched over the dish, 
and held in place by a rubber band. 

Inoculum was applied either on the upper tip or on the side of the hypocotyl. A 
1 mm square piece of brassicas cotyledon covered with conidiophores was used as 
inoculum. The Petri dish bottoms, containing the inoculated hypocotyls were placed 
in a moist chamber lined with wet paper towelling. The moist chamber was placed in 
a 15°C incubator with a light intensity of 5 ft-c for about a week. The whole hypocotyl 
was removed from solution and fixed on a slide by 0.1% cotton or anilin blue in 
lactophenol (20% carbolic acid: 20% lactic acid: 40% glycerine: 20% distilled water). 
The hypocotyl was pressed evenly with another slide until it was flattened and then 
a cover slip was applied. After a few hours, the mycelium was stained and could be 
seen within the host. The length of time between inoculation and fixation determined 
the intensity of the stain in the mycelium within the host tissue. The cotton blue 
stained the protoplasm of the fungus, but not the cell wall. Only the youngest fungal 
growth at the time of fixation was deeply stained in the final preparation. If the test 
solution favoured or did not interfere with host or fungal growth, the fungus could grow 
from the point of inoculation to the lower tip of the hypocotyl that was immersed in the 
test solution for 4 to 5 days at 15°C. At this time the mycelium in the lower tip would 
stain dark blue, but the older mycelium at the point of inoculation took little stain. If 
the test solution was unfavourable, a "zone of inhibition" lacking fungal growth could 
be measured from the base of the hypocotyl to the point where fungal growth has 
stopped. Most of the mycelium was parallel to the stele. Solutions containing either 
antibiotic or sugars were tested with this method, and for a given concentration, the 
zone of inhibition was very consistent. It was possible to use this method without 
completely aseptic procedures, and not have a serious problem with rotting. However, 
streptomycin sulfate (0.25 /ug/wl) reduced the rotting without appearing to affect host 
or the pathogen (McMeekin, 1971). 

j. Methods of breeding for multiple disease resistance 

To identify resistance to various Chinese cabbage pathogens, Williams and Leung 
(1981) developed methods for screening large populations of seedling plants. Screening 



128 

of seedlings was preferred in the early stages of breeding programs because it takes 
less time and space. Plants which exhibited seedling resistance were later evaluated 
for mature plant resistance. Such procedures may involve simultaneous inoculation 
and incubation of one week old seedlings with Plasmodiophora brassicae, Peronospora 
parasitica, Albugo Candida, Phoma lingam and Alternaria brassicicola or A brassicae. 
This can be followed by a sequential inoculation with Turnip mosaic (TUMV), or/and 
Erwinia carotovora and/or Xanthomonas campestris. The interactions phenotypes of 
more than one pathogen on a single host can be relied on for evaluation (Fig. 63) by 
paying special attention to the following: (a) careful preparation, quantification and 
delivery of precise amounts of virulent inoculum, (b) careful cultivation of host "target 
tissues" of known physiological age, and (c) optional incubation conditions for disease 
development. 

Single seeds were sown in 12-pack pots and when the cotyledons have expanded, 
after 5 to 7 days, single 0.01 to 0.02 ml drops of a freshly collected conidial suspension 
containing approximately 10 5 conidia/ml were placed on each of the two cotyledons 
with the aid of a finely tipped glass pipette (Williams and Leung, 1981). As plants of 
each 12-pack were inoculated they were placed in glass or plastic boxes containing a 
1 to 2 cm depth of warm water. A tight fitting cover was placed on the box when it was 
filled with plants. The box was then placed in a darkened incubator set at 20°C for 8- 
16h. The atmosphere in the box maintained the droplets in the cotyledons during 
which time germination and penetration occurred. Plants were then transferred to the 
greenhouse bench at 20-25°C for 5 days, then returned to the humidity boxes at 20°C 
for 16-24h. By then, the susceptible plants had a profuse growth of conidiophores on 
the lower sides of the cotyledons, whereas resistant plants exhibited varying degrees 
of sporulation and tissue necrosis which was evaluated on a 0-9 scale as illustrated in 
Fig. 62. Williams and Leung (1981) also noted that it is important to use freshly 
produced inoculum collected by washing off spores from leaves with distilled water and 
that older plants may be inoculated by atomizing the foliage with the suspension of 
conidia and keeping them at 100% RH for 8-16h. 

Chinese cabbage was grown under the above conditions and sequentially 
inoculated with four pathogens (Williams and Leung, 1981). Five days after sowing, 
seedlings were dipped in a spore suspension of Plasmodiophora brassicae spores and 
transplanted, then one to two days later inoculated with Peronospora parasitica 
conidia. Twelve days after sowing the plants were evaluated for downy mildew 
resistance and the susceptible plants removed. The remaining plants were then 
inoculated with turnip mosaic virus (TuMV) at 14 days, evaluated, reinoculated and 
rogued over the following 14 days. Surviving plants were then inoculated at 21 days 
after sowing for soft rot resistance. Two weeks later, 35 days after sowing, resistant 
plants could be removed from the pots and examined for club root. Plants 
withstanding all four diseases could then be potted and vernalized or treated with 
benlate fungicide and transplanted to the field. Further inoculations with TuMV, 



129 



TuMV 




Fig. 63. Location of inoculum placement of eight pathogens in multiple disease screening of seedling Chinese cabbage. Pb = 
P\asmodiophora brassciae, Ec = Erwinia carotovora, Ac = Albugo Candida, PI = Phoma lingam; Ab = Alternana 
brassicae; Pp = Peronospora parasitica, Xc = Xanthomonas campestris; and TUMV = Turnip mosaic virus 
(Reprinted from P.H. Williams and H. Leung. 1981. Methods for breeding for multiple disease resistant Chinese 
cabbage. ]N: Chinese cabbage. Proc. 1st Intern. Symposium, N.S. Talekar and T.D. Griggs (Editors): pp. 393-403, by 
permission of the authors and the publisher the Asian Vegetable Research and Development Center, Shanhua, Taiwan). 



130 

Erwinia and Peronospora could be made in the field and plants not treated with 
benomyl fungicide could be planted in P. brassicae infested field plots. 

The procedures for the sequential inoculations were reported to be essentially 
the same as those for individual inoculations except when TuMV inoculation was to be 
followed by E. carotovora or P. brassicae (Williams and Leung, 1981). The plants were 
maintained at 25°C instead of returning them to cooler temperatures for enhancement 
of virus symptoms. Inoculation with a combination of any of the four pathogens was 
possible by following the appropriate portions of the total sequence in Fig. 63. Plants 
can also be inoculated with other pathogens such as Albugo Candida, Alter naria spp., 
Phoma lingam and Xanthomonas campestris. Inoculation with these pathogens were 
kept apart from those areas of the cotyledons and leaves which were occupied by 
Peronospora (Fig. 64). It allowed large F 2 and backcross progenies to be efficiently 
screened. It was possible to screen approximately 600 plants per m 2 for the four 
diseases every 35 days. If resistance to each of the above pathogens were controlled 
by independent single recessive genes, a theoretical minimum population of 256 plants 
would be needed to recover the four recombinants. It is likely that far larger 
populations would be screened to accommodate the differing heritabilities for each form 
of resistance. An important consideration in selecting dominant forms of resistance in 
the production of ¥ x hybrid Chinese cabbages was that resistance to different 
pathogens can be introduced into the hybrid from different inbred parents. 

k. Heterothallism and homothalliam 

For such studies, isolates of P. parastica were collected from different host 
species and also from various geographical locations (Sherriff and Lucas, 1989b). The 
isolates were maintained on seedlings of susceptible cultivars of respective hosts. 
Cotyledons with spores were excised, placed in 50 ml sterile distilled water (SDW) and 
shaken gently to dislodge the conidia. The conidial suspension was then filtered 
through three layers of cotton gauze and centrifuged at 1500g. The conidial pellet was 
resuspended in SDW, centrifuged and finally resuspended in 1-2 ml SDW. Cotyledons 
of 7-day old susceptible seedlings, raised in a 9 cm pot, were drop inoculated with the 
conidial suspension with the aid of a Pasteur pipette. Inoculated seedlings were then 
sealed in a 13 x 21 cm propagator and transferred to a growth room (19+1°C; 16 h d, 
osram cool white fluorescent tubes, photon flux density 70/um\- 2 S" 1 ). Conidia were 
harvested 5-7 days after inoculation. 

For microscropic examination, cotyledons and leaf pieces were cleared by boiling 
for 2 minutes in a lactophenol-ethanol solution containing 10 g phenol, 20 ml glycerol, 
10 ml lactic acid and 20 ml 96% (V/V) ethanol (Sherriff and Lucas, 1989b). Cleared 
cotyledons were rinsed in water and stored in 70% (V/V) glycerol and examined under 
a low powered microscope. Mature oospores were easily visible due to their brown 



131 



DAYS 
7 14 21 

tSE£D t TRANSPLANT 



PL ASMQQfQP H QRA l 

PgRQNOSPQRA . 
PA R ASITICA . 



TuMV 

ERYV1N1A 
CAROTOVCKA 



26 



35 



25*C 





I [ 



I — [ 



too% 



RH20TC RH 20' 




INOCULATION 



OREAD INTERACTION PHENOTYPE 



Fig. 64. Sequence for individual and multiple disease resistance screening in Chinese cabbage (Reprinted from PH. Williams and 
H. Leung. 1981. Methods for breeding for multiple disease resistant Chinese cabbage. JN: Chinese cabbage. Proc. 1st 
Intern. Symposium, N.S. Talekar andT.D. Griggs (Editors): pp. 393-403, by permission of the authors and the publisher 
the Asian Vegetable Research and Development Center, Shanhua, Taiwan). 



132 

pigmentation; young oospores tended to take up and retain green pigments from host 
tissues during cleaning. 

1. Seed-borne nature of P. parasitica 

Jang and Safeeulla (1990c) studied the seed-borne nature of P. parasitica in 
Raphanus sativus. Four hundred seeds of test host cultivars were sown in field plots 
which were observed periodically for the occurrence of downy mildew disease. At the 
seed setting stage, seeds from infected plants were subjected to a maceration technique 
(Shetty et al., 1978). Seeds were placed in 250 ml of 10% NaOH for 24, 36, and 48 
hours, at 22°C along with 0.5 g of Trypan blue stain. After the alkali treatment, the 
seeds were agitated in warm water (60 - 70°C) for 5 minutes. Hard seeds were softened 
by boiling in 5% NaOH for an additional 5-10 minutes. Seeds were then sieved, 
excess water drained off, and lactophenol was added to a beaker containing the treated 
seeds. The lactophenol completed the detachment of the embryo from the seed coat. 
The beaker with the embryos and the seed coats was placed in a water bath and heated 
with a low flame until the embryos were cleared. The embryos and seed coats were 
examined under a stereomicroscope. To determine the viability of the internally borne 
mycelium, a seedling symptom test was carried out. Four hundred seeds from the 
above samples were sown under controlled conditions in a glass house which was free 
from airborne inoculum. Before sowing, the seeds were surface sterilized. Such seeds 
were sown in pots containing steam sterilized soil. Daily observations were made 
following seedling emergence and the percentage of infected seedlings within each 
cultivar was recorded. The seeds from the first harvest were subjected to the alkali 
maceration technique to determine the rate of transmission of the pathogen in the 
seeds (Jang and Sefeeulla, 1990c). 

To study pathogen infection through the stigma, unfertilized stigma of healthy 
plants were taken from test cultivars of the host (Jang and Safeeulla, 1990d). 
Unfertilized carpels were removed from healthy plants. The ovaries along with style 
and stigma were placed on the sporulating surface of infected leaves at 16°C for 3 days. 
At 12 h intervals such ovaries were fixed in acetic acid: alcohol (1:3) and subjected to 
the alkali maceration technique (Shetty et al., 1978). Another method was to spray 
unpollinated carpels with a conidial suspension, or dipping inflorescences of healthy 
plants in a container with a concentrated conidial suspension. Such treated carpels 
were covered with moist polyethylene bags to maintain humidity for 2-3 days. The 
carpels are then fixed in acetic acid: alcohol. They were dehydrated by boiling in 
alcohol: lactophenol (50:50) for 30 - 35 min. followed by maceration in 5% KOH solution 
for 24 h. The macerated carpels were washed in distilled water and treated with 
saturated chloral hydrate solution with 0.5% cotton blue for 24h. The clear ovaries 
were mounted in lactophenol on slides after squashing and then observed 
microscopically (Jang and Safeeulla, 1990d). 



133 

m. Conidial germination 

To test the effect of temperature and relative humidity on conidial germination 
and germ tube growth, a conidial suspension was made by washing off the conidia from 
the donor host leaves into petri dishes (Lin, 1981). A fine stream of cold, sterilized, 
distilled water delivered by an atomizer was used for this purpose. The donor leaves 
were usually collected at 4 a.m. when sporulation was abundant. The conidial 
suspension was adjusted to the desired concentrations (5 x 10 3 cells/ml) by dilution 
with distilled water and then sprayed on to 1.25% water agar in petri dishes. After 
incubating the inoculated petri dishes separately at 4°C, 8°C, 12°C, 16°C, 20°C, 24°C, 
28°C, 32°C and 36°C for 24h, germination and germ tube growths of conidia was 
determined by microscopic observation of 400 spores per plate. Three replications were 
used for each treatment. A conidium was considered germinated if the length of the 
germ tube exceeded the width of the conidium. To determine the effect of humidity on 
germination of conidia, three drops of conidial suspension were pipetted onto a clean 
glass slide placed in a petri dish containing saturated salt solution to obtain theoretical 
relative humidity of 0% (CaCl 2 ), 32 % (CaCl 2 6H 2 0), 55% (Ca(N0 3 ) 2 4H 2 0), 81% 
(NH 4)2 S0 4 )and 95% (NAHP0 4 12H 2 0). The petrie dishes were sealed and incubated at 
16°C for 24 h. Spore germination was then counted (Lin, 1981). 

n. Sporulation 

In the evening, diseased leaves were excised from 40 - 50 day old plants grown 
in the field (Lin, 1981). Excised leaves showing fresh symptoms were cut into several 
0.5 x 0.5 cm 2 pieces. The pieces were first gently dipped into sterilized water to wash 
off the conidia borne on conidiophores. Six pieces were then placed on a slide with the 
abaxial surface upward. The slide was put in a petri dish containing two filter papers 
previously moistened with distilled water. After incubating the petri dishes at 4°C, 
8°C, 12°C, 16°C, 20°C, 24°C, 28°C, 32°C, and 36°C, for 18 h, sporulation was 
determined by shaking the six pieces in 1 ml distilled water and counting the number 
of conidia with a haemacytometer under a microscope (Lin, 1981). 

o. Discharge of conidia 

Lin (1981) also measured the discharge of P. parasitica conidia from diseased 
leaves of the host. A leaf showing typical symptoms of downy mildew was selected and 
fixed on the hole of a spore collector so that the abaxial surface of the diseased leaf 
faced directly over one of the 24 slides attached to this collector. The surface of the 
slides were smeared with a layer of vaseline to intercept the falling conidia. Each slide 
automatically moved forward one position per hour, thus a 24 h spore collection was 
obtained. The collection was continued for three days beginning at 9 p.m. each day, 
and the periodic conidial discharge was determined by counting the conidia on slides 



134 



under a microscope. Temperature and relative humidity for each hour during conidia 
collection was also recorded to establish the relationship between the discharge of 
conidia and environmental parameters (Lin, 1981). 



13. DISEASE MANAGEMENT 

To manage downy mildew of crucifers, no single method or approach is 
considered feasible, effective, environmentally safe and economical. It is always 
essential to integrate the available methods for disease control. 

a. Cultural practices 

Cultural control of crucifers downy mildew disease is largely a matter of 
sanitation and of manipulating the environment to the advantage of the host and to 
the detriment of the pathogen. Since the pathogen survives in the form of oospores in 
the host tissues, removal, destruction and burning of the infected plant debris along 
with weeds has been suggested to restrict the source of primary inoculum (Butler, 
1918; Vasudeva, 1958). In addition, clean, well-drained soils with two years of crop 
rotation using non-cruciferous crops was also recommended. Measures to reduce the 
relative humidity around the plants by adequate aeration and avoidance of dense 
sowing and controlling the growth of weeds also helped to reduce the disease (Butler, 
1918; Conroy, 1960; Schmidt, 1960; Sherf and Macnab, 1986). Avoidance of continuous 
cropping of rape on the same field or adjacent to a field sown to rape in the previous 
year was also advised to reduce infection by P. parasitica (Downey and Bolton, 1996). 
The widespread cultivation of one or only a few cultivars of the same species may 
favour the disease. In India, the late sown crops of rapeseed-mustard were reported 
to have a higher incidence of downy mildew than the early (before October) or timely 
(by middle of October) sown crops (Kolte, 1985; Saharan, 1984, 1992a). 

In the Lujskaya area, Lenengrad regions of the USSR, the level of downy 
mildew infection was reduced on cabbage plants transplanted between 26-30 June. 
Fertilizer containing 50% humus, 45% peat and 5% millein with 3.9 g ammonium 
nitrate, 4.3g Kcl, and 8.1 g superphosphate/100 added to 110 g organic matter applied 
to the soil reduced the percentage of diseased plants better than the organic manure 
alone (Kupryanova, 1957). 

b. Seed treatment 

Fungicidal seed treatment followed by a foliar spray is a common practice to 
control downy mildew of crucifers. Metalaxyl seed treatment at the rate of 0.3 - 0.6 g 
a.i. kg' 1 reduced downy mildew infection on broccolli (Paulus and Nelson, 1977) and 



135 

rapeseed mustard (Kolte, 1985; Saharan, 1992a). A significant yield increase was 
observed when plants raised from such treated seed were sprayed once or twice with 
the same compound. Seed treatment with Apron SD 70 (35% metalaxyl and 35% 
captan) controlled downy mildew of cauliflower for more than 2 weeks after sowing 
(Crute, 1984). According to White et al. (1984) seed treatment with Apron SD 70 (1 g 
metalaxyl kg. -1 ) gave complete control of downy mildew on cauliflower inoculated 10 
days after sowing . Following seed treatment, metalaxyl was detectable in the 
cotyledons, true leaves and roots of cabbage seedlings up to 4 weeks after sowing. An 
effective and economical schedule for control of downy mildew of mustard through 
fungicidal seed treatment and/or spray application has been worked out under Indian 
conditions. Seed treatment with Apron SD 35 (2 g metalaxyl a.i. kg" 1 seed) along with 
two foliar applications of Ridomil MZ72 at 30 days intervals gave the best control of 
downy mildew on mustard along with an increase in yield (Mehta et al., 1996; Table 
30). The maximum cost-benefit ratio was obtained when mustard seeds were treated 
with Apron SD35 followed by three spays with Mancozeb. 

c. Soil treatment 

The use of systemic fungicides such as prothiocarb (Dynone) @ 5g m" 2 before 
sowing and fosetylaluminium (Aliette) @ 10 g m" 2 as soil drench gave excellent disease 
control on cauliflower (Ryan, 1977) (Table 31). Both these fungicides were as effective 
as eight sprays of dichlofluanid (Ryan, 1977). Prothiocarb also reduced infection of 
radish leaves and bulbs when applied @ 0.1% as a drench (4 litres m" 2 ) at 50% seedling 
emergence and was much more effective than sprays of dichlofluanid, zineb, captafol 
and maneb (Anonymous, 1974). 

Granular applications of metalaxyl prior to sowing was shown to be an effective 
control of downy mildew on broccoli (0.56 and 1.2 kg a.i. ha" 1 ) and on cauliflower (0.28 
kg a.i. ha" 1 *. In cauliflower, pre-sowing incorporation or a single post-sowing drench 
(1.5 kg a.i. ha" 1 ), or three high volume sprays (0.8 g a.i. litre' 1 ) of metalaxyl gave much 
better disease control than nine sprays of dichlofluanid applied during a 6-8 week 
period (Chiu, 1959). 

d. Compost treatment 

In the UK, metalaxyl, milfuran + manganese zinc dithiocarbamate, or 
propamocarb incorporated in the compost provides good control of downy mildew on 
module-raised cauliflowers in early summer plantings. In summer cauliflowers, good 
control was achieved by drenching the compost with propamocarb, fosetyl-aluminium 
foliar sprays and by applying a dichlofluanid foliar spray programme (Davies and 
Wafford, 1987). 



136 

e. Foliar spray of fungicides 

During the period from the mid-1940's to the mid-1960's, control of downy 
mildew of crucifers rested on frequent applications of sprays or dusts of fungicides such 
as chloranil (spergon), copper based materials and zineb (Channon, 1981). These 
materials were subsequently superseeded by other non-systemic fungicides like 
captafol, daconil, dichlofluanid, propineb, bordeaux mixture, copper oxychloride, 
mancozeb, ziram, chlorothalonil and fentin hydroxide (Butler, 1918; Butler and Jones, 
1949; Kolte, 1985; Saharan and Chand, 1988; Sherf and Macnab, 1986; Vasudeva, 
1958). The list of fungicides found effective against downy mildew of crucifers at 
different locations is given in Table 32. Captafol, mancozeb, difolatan, copper 
oxychloride, dichlofluanid, propineb and metalaxyl have been found to be superior to 
other fungicides on a large number of crucifers at several locations. The time of 
application of fungicides and numbers and interval of sprays depend on the duration 
and type of crop species grown (Channon et al., 1970; Kolte, 1985; Saharan and Chand, 
1988; Saharan, 1992a; Verma et al., 1994; Whitewell and Griffin, 1967). 

i) Brassica vegetables: In the UK, dichlofluanid gave excellent control of the 
disease on the cotyledons of cabbage and cauliflower. Dichlofluanid and propineb 
reduced the level of early mildew infection and increased the size and dry weight of 
cauliflower plants (Channon et al., 1970; Whitewell and Griffin, 1967). 

In the Irish Republic, downy mildew of Brassica crops, especially cauliflower, has 
been controlled by fosetyl aluminum, metalaxyl + mancozeb, cyprofuram and 
propamocarb (Ryan et al., 1984). 

In South Africa, during the initial years of containerized seedling production of 
cabbage, mancozeb (dithane M-45), chlorothalonil (Bravo), metalaxyl (Ridomil) and 
metalaxyl plus mancozeb (Ridomil MZ) provided adequate control of downy mildew 
disease. Later on, cymoxanil plus mancozeb consistently provided the most effective 
control against downy mildew. Oxadixyl plus mancozeb, cupric hydroxide and 
chlorothalonil gave significantly better protection than mancozeb (Brophy and Laing, 
1992). 

In Australia, neutralized phosphonic acid sprays applied onto cauliflower in the 
field within 3 weeks of harvest reduced downy mildew under storage conditions. Two 
applications of 2.4 kg a.i./h, 21 and 7 days before harvest reduced the curd infection 
development at the post-harvest stage in storage. There was no effect of phosphonic 
acid on crop appearance and maturity. The maximum phosphonate residue in curds at 
harvest was 12 fxg/g which was considered a safe limit (McKay et al., 1992). 

In Thailand, the best control of Chinese cabbage downy mildew was obtained with 



137 



Table 30. Efficacy, economics and spray schedule of fungicides against 
downy mildew of mustard (Mehta, Saharan and Kaushik, 1986) 



Fungicides 


Concentration 


Spray 


Percent Disease 


Percent Disease 


Percent Increase 


Cost: Benefit 




(%) 


No. 


Intensity 


Control 


In Yield 


Ratio 


Dithane M-45 


0.2 


4 


28.4 


42.4 


28.5 


1:2.20 


Kavach 


0.2 


4 


28.2 


35.0 


23.3 


1:1.57 


Ridomil MZ-72 


0.25 


3 


9.1 


81.3 


49.3 


1:1.21 


*Apron SD-35+ 














Ridonmil MZ-72 


0.25 


2 


15.8 


68.5 


34.2 


1:1.11 


Apron SD-35+ 














Dithane M-45 


0.2 


3 


22.9 


47.4 


22.1 


1:2.11 


Apron SD-35+ 














Kavach 


0.2 


3 


21.9 


49.6 


20.4 


1:1.29 


Apron SD-35 


- 


- 


33.6 


41.0 


15.9 


1:20.62 


Control 


- 


- 


49.7 


- 






LSD 0.05 


- 


- 


4.1 









* Apron SD-35 as seed treatment @ 2 g a.i. kg seed 



Ridomil 25 WP @ 2 kg/h (Yang et al., 1983). Three sprays at weekly intervals beginning 
from 28 days after transplanting gave 65% more marketable yield. 

In India, four sprays with difolatan (0.3%), daconil (0.1%), dithane M-45 (0.2%), 
Ridomil (0.2%) or aliette (0.1%) at intervals of 8-10 days were most effective for controlling 
downy mildew of radish (Sharma and Sohi, 1982; Sharma, 1983) (Table 33). Root yield was 
significantly higher in sprayed plots. There was a significant reduction in the apparent 
infection (r) and the basic infection rate (R) of downy mildew in treated plots. 

ii) Brassica oilseeds: For the control of downy mildew of mustard, difolatan, 
mancozeb and metalaxyl have been found to be very effective at different locations (Table 
34) in India. An effective and economical schedule has been worked out under Indian 
conditions for the control of downy mildew of mustard through seed treatment and/or spray 
with fungicides. Three sprays of Ridomil MZ-72 (Metalaxyl and Mancozeb @ 0.25%) at an 
interval of 20 days starting from 40 days after sowing gave maximum disease control (82%) 
along with >49% increase in yield; seed treatment with Apron SD-35 (metalaxyl @ 2 g a.i. 
kg" 1 seed) along with two foliar applications of Ridomil MZ-72 at 30 day intervals were 
relatively less effective (Tables 35, 36). These treatments were quite effective in reducing 
staghead formation in mustard (Table 37). When mancozeb (dithane M-45) and 
chlorothalonil (Kavach) were sprayed three times following seed treatment with Apron SD- 
35, disease control of around 47% and 49% respectively was achieved. The maximum cost- 



138 

Table 31. Efficacy of fungicidal treatments on the severity of downy 
mildew of cauliflower (Reprinted from E.W. Ryan. 1977. Control 
of cauliflower downy mildew (Peronospora parasitica) with 
systemic fungicides. Proc. Ninth British Insecticide and 
Fungicide Conference, Brighton, Volume 1 and 2, Research 
Report, London, U.K., Sessions 6B, pests and disease of vegetables: 
pp. 297-300, by permission of the author and the publisher British 
Crop Protection Enterprises, Loughborough, U.K.) 

Disease severity index 





Method of 
Application 


Rate of 
Application 


Walk-in 
March 28 


tunnels 
April 13 


Low tunnels 


Fungicides 


April 13 


April 25 


Dichlofluanid 


Foliar spray 


8@lg/10m 2 


1.4 


2.8 


1.6 


2.3 


Aliette 


Foliar spray 


3@lg/10m 2 


1.7 


1.9 


1.7 


1.9 


Prothiocarb 


Foliar spray 


3@lg/10m 2 


1.6 


2.1 


1.5 


1.7 


Aliette 


Soil treatment 


5g/m 2 


1.2 


2.6 


1.0 


2.0 


Aliette 


Soil treatment 


10g/m 2 


0.7 


1.8 


0.3 


1.2 


Aliette 


Soil treatment 


20g/m 2 


0.0 


0.8 


0.0 


0.6 


Prothiocarb 


Soil treatment 


5g/m 2 


0.6 


1.4 


0.2 


1.0 


Proihiocarb 


Soil treatment 


10g/m 2 


0.2 


1.1 


0.0 


0.9 


Prothiocarb 


Soil treatment 


20g/m 2 


0.0 


0.5 


0.0 


0.7 


Control 


- 


- 


2.4 


3.7 


2.0 


3.0 


LSD 5% 






0.72 


0.76 


0.78 


0.56 



O = No disease; 5 = very severe disease 



benefit ratio was either with four sprays of mancozeb or seed treatment with Apron SD- 
35, followed by three spays of mancozeb (Mehta et al., 1996). Absorption of metalaxyl 
increased, up to 30 days, when applied as seed treatment, thereafter it 
gradually declined and was not detectable after 60 days of sowing (Table 38). The 
maximum residue (average 9.03 ppm) of metalaxyl was found to be one day after 
spraying (Table 39). The metalaxyl on mustard plants was almost undetectable 15-30 
days after spraying (Table 40). The safe waiting period for metalaxyl was calculated 
to be 62 and 8 days for seed treatment and for foliar application, respectively (Table 41). 
No metalaxyl was detected in mustard seedlings raised from seeds obtained from these 
treatments (Mehta, 1993) (Table 42). 

f. Biological control 



i) Plant extracts as fungitoxicant: Garlic juice or aqueous extracts of garlic 
was reported to be toxic to P. parasitica which causes downy mildew of radish (Ark and 



139 



Table 32. Fungicides found effective against downy mildew of crucifers 



Fungicide 



Rate of Application 



Reference 



CABBAGE 

spergon spray (48% a.i.) 
spergon dust (4.8% a.i.) 
kolophygon dust (30% sulfur and 

1% phygon ) 
Parzate Dust (6.5% a.i.) 
Dithane Z-78 spray (65% a.i) 
Dithane Z-78 dust (6.5% a.i.) 
yellow cuprocide 
Dithane B-ll 
Spergon (Wittable) 
Dow Seed treatment 
Fermate 
Phygon 

Phenanthraquinone 
Bordeaux mixture 
Spergon (Chloranil) (5 and 10% a.i.) 
Dithane Z-28 (Zineb) (3.9% a.i.) 
Phygon XL(dichlone) (1.0% a.i.) 
Copper No. 30 (4.0% a.i.) 
Thiram (5.0% a.i.) 
Manzate (Maneb) (4.2% a.i.) 
Vancide F995W (6.0% a.i.) 
Vancide 51ZW (6.0% a.i.) 
Ethyl B-622 (4.0% a.i.) 
Metalaxyl 

Captafol (0.25% a.i.) 
Daconil 2787 
Dichlofluanid 
Propineb 
Zineb 
Maneb 
Mancozeb 
Quintozene 
Quinomethionate 
Copper oxychloride 
Triphenyl tin hydroxide 
Dichlone 
Chloranil 

Nabam - Zinc Sulphate 
Phygon XL-N 
Difolatan 4F (Captafol) 
Polyram (Metiram) 
Dithane Z-78 
Dichlofluanid 



4 lbs/100 gallons 
30 lbs/acre 

30 lbs/acre 

30 lbs/acre 

2 lbs/100 gallons 

30 lbs/acre 

1 lb/100 gallons 

1 lb/100 gallons 
4 lbs/100 gallons 

2 lbs/100 gallons 
2 lbs/100 gallons 
1/4 lb/100 gallons 
1 lb/100 gallons 
1:1:10 



Borders, 1953 



Foster 1947 b 



Anonymous, 1938, Wiese, 1927 
Epps, 1955 



1.12kga.i./ha 



Jaworski et al., 1982 
Channon & Hampson,1968 



4 lbs. 48%/100 gallons 

1 lb/100 gallons 

1 lb/100 gallons 

0.2% 

0.2% 

0.2% 

0.05-0.2% a.i. 



Anonymous, 1953 



Apandi, 1980 

Ciferri, 1953 
Channon et al., 1970 



140 



Aspor 

Maneb 

Perotsin 

Nickel sulphate spray 

Polycarbacin spray 

Cymoxanil + mancozeb (6+70% a.i.) 

Cymoxanil + chlorothalonil (6+50% a.i.) 

Oxadixyl (8% a.i.) 

Oxadixyl + mancozeb (8%+56% a.i.) 

Propamocarb + HC1 (72% a.i.) 

Propamocarb + mancozeb (72 + 80% a.i.) 

Metalaxyl (Ridomil WP) (25% a.i.) 
Metalaxyl + Mancozeb (25 + 80% a.i.) 
Fosetyl - Al - Mancozeb (44 + 26% a.i.) 
Chlorothalonil (50% a.i.) 
Mancozeb (80% a.i.) 
Copper oxychloride (80% a.i.) 
Cupric hydroxide (72% a.i.) 
CGA - 48988 soil application 



0.3% 
0.2% 
0.3% 

0.05-0.2% 
0.4% 

200g/100 litres 
200g/100 litres 
80ml/100 litres 
330g/100 litres 
120ml/100 litres 
60ml/100 litres+ 
75g/100 litres 
50g/100 litres 
50g/100 Utres 
350g/100 htres 
lOOml/100 htres 
200g/100 litres 
400g/100 litres 
200g/100 litres 
23 mg a.i./M 



Nakov, 1968 



Keyworth, 1967 
Vasileva, 1976 
Brophy & Laing, 1992 



Gabrielson & Getzin, 1979 



BROCCOLI 

Agrimycin 

Spergon SL 

Streptomycin 

Agri-strep 

Agri-strep + Glycerol 

Copper-Zinc 

Copper-Manganese 

Spergon SL 

Manzate 

Thioneb 

Captan 50W 

Vancide M 

Kemate 50% 

Manzate + Agri-strep 

Copper-zinc + Agri-strep 

Agrimycin 500 

CGA- 1-82 50 WP 

CGA-38140 50WP 

CGA-48988 (Metaxadine) 



0.1 lb/acre 

2 lbs/acre 
50 ppm 

3 lbs/acre 
3 lbs/acre 
6 lbs/acre 
6 lbs/acre 

3 lbs/acre 

4 lbs/acre 
6 lbs/acre 
6 lbs/acre 
4 lbs/acre 
6 lbs/acre 
4+0.4 lbs/acre 
4+0.4 lbs/acre 
4.6 lbs/acre 

2 lbs/5 ft. band 
soil application 
2 oz/acre 14 days 
after seeding 
1 or 2 oz/100 lbs seed 



Natti et al., 1956 

Altaian, 1958 
Natti, 1957 



Natti, 1957, 1959 
Natti, 1957 

Johnston & Springer, 1977 



Paulus et al., 1978 



CAULIFLOWER 

Dichlofluanid 50WP 
Captafol 85WP 
Zineb 70WP 
Propineb 70WP 
Daconil 2787 75WP 



1 V2 lbs/100 gallons 
3 lbs/100 gallons 
3 lbs/100 gallons 
3 lbs/100 gallons 
3 lbs/100 gallons 



Whitewell & Griffin, 1967 



141 



Dichlofluanid 






0.05-0.2% a.i. 


Fosetyl aluminium 






- 


Metalaxyl + Mancozeb 






- 


Cyprofuram 






- 


Propamocarb 






- 


Phosphonic acid 






2.4 kg a.iVh 


RAPESEED-MUSTARD 








Polyram M 






2 lbs/100 gallons 


Melprex 






1.5 lbs/100 gallons 


Bordeaux mixture 






4:4:50 (0.8%) 


Cuprovit 






2 lbs/100 gallons 


Dithane M-45 






2 lbs/100 gallons 


Dithane M-45 






0.3% 


Dithane Z-78 






0.3% 


Blitox-50 






0.3% 


Difolatan 80 






0.2% 


Ziram 






0.2% 


Dithane M-45 






0.2% 


Thiovit 






0.2% 


Difolatan 






0.2% 


Dithane M-45 






0.2% 


Dithane Z-78 






0.2% 


Blitox 50 






0.3% 


Ridomil 






0.2% 


Bristan 






0.1% 


Apron SD-35 






0.2% seed treatmei 


Metalaxyl 






0.2% 


Kavach 






0.2% 


Radomil MZ-72 






0.25% 


Apron SD 35 seed treatment 






2 g a.i./kg seed 


+ Dithane M-45 spray 






0.2% 


Apron SD 35 seed treatment 






2 g a.i./kg seed 


+Radomil MZ-72 spray 






0.2% 


Apron SD 35 seed treatment 






2 g a.i./kg seed 


+ Difolatan spray 






0.2% 


Apron SD 35 seed treatment 






2 g a.i./kg seed 


+ Kovach spray 






0.2% 


RADISH 








Difolatan seed treatment 


or 


spray 


0.3% 


Daconil seed treatment 


or 


spray 


0.1% 


Dithane M-45 seed treatment 


or 


spray 


0.2% 


Ridomil seed treatment 


or 


spray 


0.1% 


Aliette seed treatment 


or 


spray 


0.1% 


Blitox seed treatment 


or 


spray 


0.2% 


Captan seed treatment 


or 


spray 


- 


Copper oxinate seed treatment 


or 


spray 


- 


Delan seed treatment 


or 


spray 


- 


Dathane Z-78 seed treatment 


or 


spray 


- 


Macuprax seed treatment 


or 


spray 


- 



Channon et al., 1970 
Ryan et al., 1984 



McKay et al., 1992 



Perwaiz et al., 1969 



Bains & Jhooty, 1979 



Chauhan & Muheet, 1976 



Saharan, 1984; 1992a 



Mehta et al., 1996 



Sharma & Sohi, 1982 



142 

STOCK 

Zineb 8 lbs/100 gallons Jafar, 1963 

Strepto spray + glycerol 500 ppm + 1% 

Trioneb 8 lbs/100 gallons 

Bordeaux mixture 5:5:50 

Fongarid (CGA 38140) 0.05% Trimboli & Hampshire, 1978 

Zineb 0.13% 

CAMELINA SATrVA 

Brestan - Zarzycka & Kloczowska, 1964 

Polyram-M 

Sadoplon 

Copper oxychloride 0.4% Zarzycka & Kloczowska, 1967 



Thompson, 1959). 

ii) Antagonists for biocontrol: Bacteria were observed on the mycelium, 
conidiophores and conidia of P. parasitica on Lepidium graminifolium (Nicolas and 
Aggery, 1940). This was associated with a reduction in conidial germination. 

g. Host resistance 

Many sources of resistance to downy mildew of crucifers have been identified in 
major host species from various parts of the world. Information is also known on the 
genetics of the host parasite interaction. Efforts are being made to breed downy mildew 
resistant cultivars in various crucifers through conventional and biotechnological 
techniques. 

h. Integrated disease management 

In the quadrangle of integrated control (chemical-cultural-biological-host 
resistance) of downy mildew of crucifers, biological control has not been exploited at the 
field scale. Breeding for resistance has only succeeded in some crucifers. Chemical 
control of the disease may not always be reliable as resistance has been developed in 
P. parasitica to metalaxyl, which at one stage proved outstandingly effective in the 
control of downy mildew (Brophy and Laing, 1992; Crute et al., 1985). Thus there is 
clearly a need to breed sources of host resistance that would counter pathogenic 
variation. It is also possible that differential sources of host resistance could be useful 
in programs of integrated control if they were deployed together with fungicides; this 
would potentially prolong the effectiveness of both control procedures (Silue et al., 
1996). Other methods involve sanitation, field practices like sowing time, plant density, 
and the judicious use of nutrition and irrigation so that inoculum levels will not build 



143 



Table 33. Efficacy of fungicidal sprays on downy mildew of radish 
(Reprinted from S.R. Sharma and H.S. Sohi. 1982. Effect of 
fungicides on the development of downy mildew and white rust 
of radish. Indian J. Agric. Sci. 52:521-524, by permission of the 
authors and the publisher Indian Council of Agricultural 
Research, New Delhi, India) 









Apparent infection 


Basic infection 






Fungicides 


Disease index (%) 
A B 


rate (r) 


rate (R) 




Yield of roots 
A 


; (ke/dot) 




A 


B 


A 


B 


B 


Aliette 


NT 


6.37 


NT 


0.078 


NT 


0.59 


NT 


20.45 


Blitox 


17.07 


12.27 


0.108 


0.088 


2.36 


0.93 


18.45 


22.02 


Captan 


17.18 


11.03 


0.115 


0.083 


3.08 


0.79 


19.85 


19.65 


Copper oxinate 


24.46 


8.25 


0.128 


0.083 


4.60 


0.81 


18.85 


18.47 


Deconil 


3.70 


3.40 


0.055 


0.051 


0.28 


0.21 


23.15 


24.45 


Delan 


19.75 


8.22 


0.123 


0.083 


4.04 


0.81 


19.67 


19.17 


Difolatan 


5.88 


4.10 


0.079 


0.053 


0.81 


0.23 


25.67 


24.06 


Dithane M-45 


7.35 


7.55 


0.084 


0.071 


0.99 


0.48 


22.37 


24.12 


Dithane Z-78 


13.67 


10.30 


0.109 


0.092 


2.56 


1.10 


21.52 


21.61 


Macuprax 


26.82 


13.20 


0.130 


0.101 


4.84 


1.53 


17.92 


18.43 


Control 


55.79 


19.51 


0.171 


0.109 


13.06 


1.93 


17.30 


17.73 


SEM+ 


1.74 


0.942 










1.156 


1.020 


CD at 5% 


5.049 


2.722 










3.356 


2.947 



A = December 1979 - February 1980; B = June - September 1980; NT = Not tested 



Table 34. 


Efficacy of 


fungicidal treatments 


on the downy 


mildew of 




mustard in 


India (Saharan, 


1984, 


1992a) 






Concentration % 




Percent Disease Intensity 


Fungicide 


Durgapura 


Hisar 


Pantnagar 


Defolatan 


0.2 




10.65 




9.80 


15.09 


Dithane M-45 0.2 




8.31 




12.00 


10.79 


Dithane Z-78 


0.2 




- 




14.80 


14.50 


Blitox 


0.2 




15.00 




14.80 


16.77 


Ridomil 


0.2 




16.25 




8.0 


- 


Control 


- 




23.00 




24.80 


16.80 


CD. 5% 


- 




- 




5.90 


5.69 



144 



Table 35. 



Efficacy and spray schedule of fungicides against downy mildew 
of mustard during 1991-92 and 1992-93 crop seasons (Mehata, 
Saharan and Kaushik, 1996) 























+ Percent 




Cone. 

% 


No. of 
Sprays 




+ 


Percent Disease Index (DAS) 


Disease Control 


Treatments 


1991-92 


1992-93 


Averaee 


1991-92 
(60) 


1992-93 
(90) 


Averaee 




(60) 


(90) 


(60) 


(90) 


(60) 


(90) 




Dithane M-45 


0.2 


4 


2.7 


26.2 


11.3 


30.7 


7.0 


28.4 


39.7 


45.2 


42.4** 


Kavach 


0.2 


4 


3.5 


28.2 


- 


- 


3.5 


28.2 


35.0 


- 


35.0 


Difolatan 


0.2 


4 


2.7 


22.4 


- 


- 


2.7 


22.4 


48.2 


- 


48.2 


Ridomil MZ-72 


0.25 


3 


0.2 


7.8 


4.7 


10.3 


4.0 


9.1 


82.0 


81.6 


81.8** 


*Apron SD-35+ 


$ 






















Ridomil MZ-72 


0.25 


2 


0.0 


12.9 


12.3 


18.7 


6.1 


15.8 


70.3 


66.6 


68.5** 


*Apron SD-35+ 
























Dithane M-45 


0.2 


3 


0.4 


22.9 


- 


- 


0.4 


22.9 


47.4 


- 


47.4 


*Apron SD-35+ 


0.2 


3 


1.2 


21.9 


- 


- 


1.2 


21.9 


49.6 


- 


49.6 


Kavach 
























* Apron SD-35+ 


0.2 


3 


1.8 


20.9 


- 


- 


1.8 


20.9 


51.8 


- 


51.8 


Difolatan 
























* Apron SD-35 


*. 


- 


2.2 


27.3 


15.3 


40.0 


8.7 


33.6 


36.9 


28.6 


32.7** 


Unsprayed 


- 


- 


4.8 


43.4 


23.7 


56.0 


14.2 


49.7 


- 


- 


- 


(control) 

























+ Average of four replicates 
*Seed treatment @ 2. g a.i. kg' 1 seed 



( ) DAS: Days after sowing 
**: Two years mean 



up too rapidly. 

Butler (1918) reported a long time ago that the disease can be controlled in young 
crucifer plants by a mulch of sawdust saturated with copper sulphate placed around 
the base of the plants. 

In the Shanghai region of China, a combination of seed treatments, direct 
seeding, application of fertilizer, and 2-3 fungicide sprays at the first peak infection 
period decreases the incidence of P. parasitica in Chinese cabbage and increases yield 
by 10 - 18% (Shao et al., 1991). 



145 

14. FUTURE STRATEGIES AND PRIORITIES OF DOWNY MILDEW 
DISEASE MANAGEMENT 

With the globalization of agriculture, there is significant increase in the 
movement of crucifer germplasm and in the cropping patterns of this important 
commodity all over the world. The absence of strict measures on the restriction of 
movement of germplasm and the intensive cultivation of these crops has resulted in 
large scale perpetuation, build-up and dissemination of Peronospora parasitica 
virulences on cruciferous species all over the world. The information gathered in this 
monograph indicates that some gaps still exist in the complete comprehension of this 
disease and this is indicated below: 

a. Disease epidemiology 

Factors governing disease initiation, development and consequent progression are 
not completely understood. There is need for more information on the role of initial 
inoculum in disease development, in the area of changing host susceptibility over time 
and in oosporic multiplication. Time should always be included as one of the variables 
in the study of the relationship between pathogen development and host or 
environmental conditions. Multilocational trials with staggered dates of planting can 
be helpful in analyzing disease development in relation to environmental conditions 
and to develop disease prediction models. 

b. Physiological specialization 

To analyze the virulence pattern of P. parasitica, identification and 
standardization of host differentials is necessary. The relationship between 
pathogenicity on wild hosts and crop plants needs further study since wild hosts may 
act as a theatre for increased genetic variation in the pathogen. The use of modern 
techniques like RAPD fingerprinting may distinguish between pathotypes or even 
separate clonal population. PCR amplification of ITS (internal transcribed spacer) 
regions may be useful both for identification of isolates and in estimating their 
similarity. Ultimately this type of sequence analysis may reveal the evolutionary 
relationship between different species, genera or higher taxa. The studies on variation 
between isolates obtained from different geographic regions might explain the 
variability with respect to virulence and to other characters such as fungicide 
sensitivity. 

c. Genetics of resistance 

The search for new sources of resistance is always of high priority. The 
understanding of population biology and genetics (genetic diversity, relative fitness in 
geographically separated population, good knowledge of host pathogen variation, 
availability of reliable markers like virulence/avirulence mating type, allozymes, 



146 



Table 36. Comparative yield increase and cost benefit ratio of fungicides 
used against downy mildew of mustard (Mehta, Saharan and 
Kaushik, 1996) 



Treatments 


♦Average 


% increase 


Cost: benefit** 




yield/ 


in yield 


ratio 




plot* (kg) 


over control 


Rs.: Rs. Ps. 


Dithane M-45 


1.164 


28.5 


1:2.20 


Kavach 


1.116 


23.3 


1: 1.57 


Ridomil MZ-72 


1.352 


49.3 


1:1.21 


***Apron SD-35 + 


1.216 


34.2 


1:1.11 


Ridomil MZ-72 








Apron SD-35+ 


1.115 


22.1 


1:2.11 


Dithane M-45 








Apron SD-35+ 


1.090 


20.4 


1:1.29 


Kavach 








Apron SD-35 


1.050 


15.9 


1:20.62 


Unsprayed 


0.950 


- 


- 


(control) 









+ 
* 

** 



Average of four replicates 

Plot size: 2.0 x 2.1m 2 

Based on prelavent market price in 1992 



*** 



Raya: Rs.800/-Q 

Dithane M-45: 152/kg 

Ridomil MZ-72:950/kg 

Kavach: 333/kg 

Apron SD-35: 2782/kg 

Labour - 5 labour/spray/hectare @ Rs.40/ - per labour 

Seed treatment @ 2 g a.i. kg" 1 seed 



mitochondrial and nuclear DNA RFLPS) should lead to a more effective management 
strategy for disease control. 



On the cellular level in relationship to molecular studies, the development of an 
axenic culture system would help in studies of heredity. Different resistance 



147 



Table 37. Efficacy of fungicides against staghead of mustard due to 
combined infection of white rust and downy mildew (Mehta, 
Saharan and Kaushik, 1996) 



Treatments 


No.of 


+ Staghead** 


+ Staghead** 


+ Staghead** 




Sprays 


incidence (%) 


length (cm) 


score 


Diathane M-45 


4 


8.8 


(16.2) 


7.9 


2.1 


Kavach 


4 


2.4 


(8.9) 


8.9 


1.4 


Ridomil MZ-72 


3 


1.6 


(6.4) 


5.4 


1.5 


*Apron SD-35 + 


2 


5.2 


(12.0) 


8.1 


2.0 


Ridomil MZ-72 












*Apron SD-35+ 












Dithane M-45 


3 


2.6 


(9.2) 


1.9 


1.6 


*Apron SD-35+ 












Kavach 


3 


2.9 


(9.9) 


3.7 


1.7 


* Apron SD-35 


- 


13.9 


(19.0) 


10.4 


2.3 


Unsprayed 


- 


25.8 


(30.4) 


14.5 


3.4 


(control) 












LSD (0.05) 






(2.7) 







() 

+ 
* 

** 



Angular transformed values 
Average of four replicates 
Seed treatment @ 2 g a.i.kg" 1 seed 
Two years mean (91-92, 92-93) 



mechanisms should be characterized in more detail at the histological and cellular 
level. 



d. Molecular aspects 

On the molecular level, a major emphasis should be placed on the development and 
improvement of methods for isolation of RNA and DNA, isozyme analysis, use of RFLP 
analysis for assessment of genetic variation, development of genetic maps, research of 
transposable elements and plasmids, development of fungal transformation system, i.e., 
availability of vectors with suitable markers, and methods for introducing DNA. The 



148 



Table 38. Persistence of metalaxyl in mustard foliage after seed treatment 
(Mehta, 1993) 



Treatment 


Days of 


Average 


* Range 


Dissipation 


SD± 




sampling 


residue 
Level (ppm) 


(ppm) 


(%) 




Apron SD-35 


7 


1.81 


1.58-2.02 


0.00 


0.23 


@ 2 g a.iAg seed 


15 


3.46 


3.03-3.80 


+191.46 


0.37 




30 


9.08 


8.63-9.84 


+242.42 


0.54 




40 


5.82 


5.57-6.01 


35.90 


0.18 




60 


0.00 


- 


100.00 


- 



Average of three replicates. 



Table 39. Persistence of metalaxyl in foliage of mustard after foliar 
application (Mehta, 1993) 



Treatments 



Average residue level* (ppm) 





Days 


1 


5 


10 


15 


30 


** Foliar spray-I 




9.03 


0.68 


0.28 


0.0 


0.0 


Range 




7.94-10.08 


0.49-0.80 


0.25-0.32 


- 


- 


Dissipation (%) 




0.00 


92.46 


96.89 


100.0 


100.0 


Foliar spray - II 




10.37 


0.54 


0.29 


0.0 


0.0 


Range 




9.57-11.10 


0.51-0.57 


0.29-0.30 


- 


- 


Dissipation (%) 




0.00 


94.79 


97.20 


100.0 


100.0 



SD+ I Spray 
II Spray 



0.830 
0.812 



0.169 
0.034 



0.040 
0.005 



** 



Average of three replicates 

Fnlinr snmv: 40, 70 days after sowing @ 0.25% 



Foliar spray: 



149 



Table 40. Persistence of metalaxyl in mustard foliage after seed treatment 
and foliar sprays (Mehta, 1993) 



Treatments 



Average residue level* (ppm) 





Days 1 


5 


10 


15 


30 


** Foliar spray- 1 


8.21 


0.49 


0.26 


0.0 


0.0 


Range 


7.89-8.76 


0.47-0.52 


0.23-0.29 






Dissipation (%) 


0.00 


94.03 


96.83 


100.0 


100.0 


Foliar spray - II 


9.45 


0.69 


0.27 


0.0 


0.0 


Range 


8.64-10.22 


0.59-0.80 


0.25-0.30 


- 


- 


Dissipation (%) 


0.00 


92.69 


97.14 


100.0 


100.0 



SD± I Spray 
II Spray 



0.382 
0.673 



0.028 
0.121 



0.034 
0.029 



** 



Foliar sprays : 60 & 80 days after sowing @ 0.25% 
Average of three replicates. 



Table 41. Safe period and residue half life values of Metalxyl in mustard 
(Mehata, 1993) 



Treatment 



RL-50 

(Days) 



SWP 

(Days) 



Seed treatment 
Foliar Spray 1 
Foliar Spray 2 



17.57 

2.08 

1.79 



62.33 
8.69 
7.85 



RL-50 = Residue half life 
SWP = Safe waiting period 



150 



Table 42. Translocation of metalaxyl residues into mustard seed following 
different treatments at harvest (Mehta, 1993) 



Treatment Average residue (ppm) 



Seed-treatment* (Apron SD-35) (T x ) ND 

60 DAS 

Foliar sprays** Ridomil MZ-72) (T 2 ) ND 

40, 70 DAS 

Seed treatment + Foliar spray (T 3 ) ND 

60, 80 DAS 



ND: Not Detectable DAS: Days after sowing 

* @ 2 g a.i. kg-1 seed ** @ 0.25 per cent 



cloning of avirulence and resistance genes will contribute to the elucidation of the 
molecular basis of host-pathogen specificity and from the practical point of view, be 
helpful in the design of integrated control strategies. 

The concentrated effort to identify loci in Arabidopsis thaliana associated with 
specific resistance to P. parasitica should enable cloning of host resistance genes in the 
near future. The eventual isolation and functional analysis of avirulence genes from 
crucifers downy mildew will be of particular interest, given the apparently separate 
phylogeny of this group from other plant pathogenic fungi. 

e. Biochemical aspects of resistance 

The elucidation of the biochemical background of biotrophy, the establishment of 
an intracellular interface with host cells, and the role of different infection structures 
should be topics of future research. Also we understand very little of the biochemical 
mechanisms involved in the hypersensitive reaction and in various types of resistance. 
There is a need to gather information concerning the effects of the downy mildew 
fungus on respiration, photosynthesis, and the translocation, accumulation and transfer 
of carbohydrates in infected host tissues. The role of hormonal disturbances in 
pathogenesis, and the basis of systemic versus local lesion infection needs more study. 



151 

Genetical and histo-cytological descriptions of interactions, and the availability of 
methods for growing parasites alone and in combination with their hosts, are largely 
lacking. 

f. Disease management 

There is good information on the efficacy of fungicides against downy mildew 
pathogen. Efforts should continue to search for low cost effective chemicals which can 
provide economically significant disease control. The possibility of biocontrol agents 
need to be explored. Study of integrated disease control strategies may be very useful. 
However, integration of all the means of control needs to be done for each crop and for 
each geographical region. 



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159. 



180 
16. SUBJECT INDEX 
Abiotic Elicitors: 110. 
Aggressive: 57. 

Albugo candidal 5, 11, 69, 73, 74, 76, 83, 87-91, 128. 
Allyl isothiocyanate Effect: 109, 110. 
Antibiotics: 140. 

Agrimycin: 140. 

Agri-strep: 140. 

Streptomycin: 120, 140, 142. 
Appressoria: 25, 47, 48, 73, 76. 
Arabidopsis species: 51, 59, 106, 108. 
Area Under Disease Progress Curve: 83, 84, 89. 
Asexual Phase: 20. 
Axenic culture: 64, 65. 

Biochemical Compounds: 92-99, 103, 110, 150. 
Biochemical Basis of Resistance: 109, 110, 150. 
Biological Control: 138, 142. 
Botrytis parasitica: 20. 
Brassica Oilseeds: 5, 9, 57, 83, 88, 137. 
Brassica Vegetables: 5, 15, 88, 136. 
Brassica: 9, 14, 15, 56-60, 88. 

B. alba: 14, 51, 55-58, 117. 



181 
B. alboglabra: 6. 

B. campestris: 14, 51, 55-60, 64, 69, 108, 114, 117.. 

B. carinata: 14, 56-59, 117. 

B. caulorapa: 57, 58. 

B. chinensis (Chinese Cabbage): 6, 9, 55-58, 69, 83, 118, 128, 129, 131, 136, 144. 

B. fructiculosa: 51, 55. 

B.juncea: 5, 7, 9, 11, 51, 55-51819, 62, 69, 84-91, 98, 108, 110, 111, 114, 117. 

B.juncea var. megarrhiza (Chinese Mustard): 7, 9, 56. 

B.juncea var. multiceps (Chinese Mustard): 6, 7, 9, 56. 

/?. napobrassica: 55. 

5. na/ws: 6-8, 51, 55-62, 99, 108, 110, 114, 118. 

B. nigra: 6-8, 51, 56-59, 118. 

B. oleracea var. botrytis (Broccoli): 6, 7, 9, 13, 16, 56, 88, 106, 107, 114, 117, 135, 146. 

B. oleracea var. botrytis (Cauliflower): 6- 9, 13, 20, 23, 56-59, 69, 73, 83, 99-104, 
109, 114, 117, 126, 135, 136, 138. 

B. oleracea var. capitata (Cabbage): 5-9, 13, 23, 26-28, 51, 55-60, 63, 72, 74, 
76, 79-83, 88, 93-97, 99, 107, 109, 114, 117, 119, 136, 139, 140. 

B. oleracea var. gemmifera (Brussel's Sprouts): 6, 7, 9, 114. 

B. oleracea var. gongylodes (Kohlrabi): 6-9, 13, 65, 73, 114. 

B. oleracea var. viridis (Kale): 6, 7, 9, 114. 

B. pekinensis (Chinese Cabbage): 6, 7, 9, 55-58, 63, 65, 66, 69, 71-73, 107, 123. 

B. rapa: 5-7, 9, 11, 51, 55-59, 69, 108, 118. 

B. tournefortii: 51, 57-59. 



182 
Brassica species: 5-9, 20, 24, 51, 55-60, 100, 101, 125. 

Brassica species: (Rapid Cycling): 58, 59, 64, 108. 

Breeding for Disease Resistance: 114, 115, 127, 128, 131. 

Callus Culture: 119, 120. 

Camelina species: 6, 142. 

Camelina sativa: 6, 8, 142. 

Capsella bursa-pastoris: 6-8, 20, 55, 56, 59. 

Cardamine impatiens: 7. 

Caronopus didymus: 8. 

Chemical Dust: 136. 

Chemical Spray: 136-144, 146-150. 

Colza: 6, 8, 9. 

Compositae: 4. 

Compost Treatment: 135. 

Conidial Discharge: 65-68, 133. 

Conidial Germination: 24, 60, 69-73, 79, 133. 

Conidial Longevity: 63, 64. 

Conidial Measurement: 22-24, 56, 59. 

Conidiophore and Conidia: 24, 41, 45, 49, 50, 64, 65, 72, 83. 

Conidiophore Development: 38, 40-45, 49, 54. 

Control: 134-151. 

Cost-Benefit Ratio: 137. 



183 
Crop Rotation: 134. 

Crucifers: 4-10, 13-15, 17, 18, 20, 51, 56, 64, 74, 76-78, 98, 99, 136, 139. 

Cultivar: 13, 57, 62, 64, 65, 67, 85, 90, 99, 107-109, 111, 117, 118, 125. 

Culture Medium: 64, 119, 120. 

Cultural Practices: 134. 

Cytology and Genetics: 45, 63. 

Detached Leaf (Cotyledon) Culture: 63, 120, 121, 124, 125. 

Differential Resistance: 57, 58, 60, 64, 108, 109, 114. 

Disease: 4. 

Disease Assessment: 17, 19, 122. 

Disease Cycle: 74, 76, 77. 

Disease Development: 76, 84, 89-91. 

Disease Incidence/Intensity/Severity: 17, 18, 88-91, 93, 98, 122, 137, 138, 143. 

Disease Index: 18, 89-91, 110, 137, 143, 144. 

Disease Intensity - yield loss equation: 5, 9. 

Disease Management: 134-151. 

Disease Occurrence: 6-8, 88. 

Disease Perpetuation: 74. 

Disease Rating: 17, 122. 

Disease Scoring Scale: 17, 18, 122, 123. 

Disease Transmission: 74. 

Downy Mildew: 4, 5, 8, 9, 11, 17, 18, 20, 45, 51, 74, 78, 83-91, 98, 99, 111, 122, 135-137, 139, 



184 
143, 144, 146. 
Economic Importance: 5. 
Electron Micrograph: 26-28, 30-37, 47-50. 
Electron Microscopy: 25-28, 30-38, 45, 47-50. 
Enzymes: 74, 93, 99, 103, 104, 113, 114. 
Epidemiology: 76, 145. 
Eruca sativa (Taramira): 6, 7, 9, 51, 56, 57. 
Erucic Acid: 110. 
Fertilizer Effect: 88, 134, 144. 
Fungicide Residue: 138, 148-150. 
Fungicide Resistance: 115, 116, 119, 126. 
Fungicide Spray: 136-144, 146-150. 
Fungicide Tested: 125, 126, 139-142. 

Aliette: 135, 137, 138, 141, 143. 

Apron: 135, 137, 138, 141, 144, 146-148. 

Aspor: 140. 

Blitox 50: 141, 143. 

Bordeaux mixture: 136, 139, 141, 142. 

Bravo: 136. 

Brestan: 141, 142. 

Captafol: 135, 136, 139, 140. 

Captan 50W: 140, 141, 143. 



185 



CGA 1-82, 38140, 48988: 140. 

Chloranil: 136, 139. 

Chlorothalonil: 136, 140. 

Copper Oxide: 136. 

Copper Oxinate: 141, 143. 

Copper Oxychlorjde: 136, 139, 140, 142. 

Copper Sulphate: 144. 

Cupravit: 141. 

Cupric Hydroxide: 136, 140. 

Cuprocide: 139. 

Cymoxanil: 136, 140. 

Cyprofuram: 136, 141. 

Daconil: 136, 137, 139-141, 143. 

Delan: 141, 143. 

Dichlofluanid: 135, 136, 138-141. 

Dichlone: 139. 

Difolatan: 136, 137, 139, 141, 143, 144. 

Dithane B-ll: 139. 

Dithane M-45: 136, 137, 141, 143, 144, 146, 147. 

Dithane Z-28: 139. 

Dithane Z-78: 139, 141, 143. 

Dow Seed Treatment: 139. 



186 
Dynone: 135. 
Ethyl B-622: 139. 
Fentin Hydroxide: 136. 
Fermate: 139. 
Fongarid: 142. 

Fosetyl aluminium: 135, 136, 140, 141. 
Kavach: 137, 141, 144, 146, 147. 
Kemate: 140. 
Kolophygon: 139. 
Macuprax: 141, 143. 
Mancozeb: 135, 136, 138-141. 
Maneb: 135, 139, 140. 
Manzate: 135, 139, 140. 
Melprex: 141. 

Metalaxyl: 115, 116, 119, 126, 134-138, 140-150. 
Metaxadine: 140. 
Milfuran: 135. 
Nebam-Zinc Sulphate: 139. 
Nickel Sulphate: 140. 
Oxadyxil: 136, 140. 
Parzate: 139. 
Perotsin: 140. 



187 
Phenanthraquinone: 139. 

Phenylamide: 58 

Phosphonic Acid: 136, 141. 

Phygon: 139. 

Polyram M (Metiram): 139, 141, 142. 

Polycarbacin: 140. 

Propamocarb: 135, 136, 140, 141. 

Propineb: 136, 139, 140. 

Prothiocarb: 135, 138. 

Quintomethionate: 139. 

Quintozene: 139. 

Ridomil: 115, 116, 119, 126, 134-150. 

Sadoplon: 142. 

Spergon: 136, 139, 140. 

Sulphur: 139. 

Thiovit: 141. 

Thiram: 139. 

Thioneb: 140. 

Trioneb: 142. 

Triphenyl Tin Hydroxide: 139. 

Vancide: 139, 140. 

Zineb: 135, 136, 139, 140. 



188 
Ziram: 136, 139, 141, 142. 

Garlic Juice: 138. 

Genetics of Host-Pathogen Relationship: 106-109, 145. 

Geographic Distribution: 5, 6. 

Argentina: 6. 

Australia: 6, 7, 136. 

Austria: 6. 

Bavaria: 6. 

Bermuda: 6. 

Borneo: 6, 

Brazil: 6. 

Brunei: 6. 

Bulgaria: 6, 57. 

Canada: 6. 

Chile: 6. 

China: 6, 144. 

Costa Rica: 6. 

Cuba: 6. 

Cypress: 6. 

Czechoslovakia: 6. 

Denmark: 6. 

Dominica: 6. 



189 
Ethiopia: 6. 

Fiji: 6. 

Finland: 6. 

France: 6, 57. 

Germany: 6. 

Greece: 6. r 

Guatemala: 6. 

Haiti: 6. 

Holland: 6, 51. 

Hong Kong: 6. 

Hungary: 7. 

Iberica (Spain): 7. 

India: 7, 57, 58, 83, 134, 143. 

Iran: 7. 

Iraq: 7. 

Ireland: 7, 136. 

Israel: 7. 

Italy: 7. 

Jamaica: 7. 

Japan: 7. 

Kampuchea: 7. 

Kenya: 7. 



190 
Korea: 7. 
Libya: 7. 
Ludlow: 7. 
Malawi: 7. 
Malaysia: 7. 
Malta: 7. 
Mauritius: 7. 
Mexico: 7. 
Montpellier: 7. 
Moravea: 7. 
Morocco: 7. 
Mozambique: 7. 
Nepal: 7. 
Netherlands: 7. 
New South Wales: 7. 
New Zealand: 7. 
North Borneo: 6. 
Norway: 7. 
Pakistan: 5, 7. 
Palestine: 7. 
Panama: 7. 
Papua New Guinea: 7. 



191 
Philippines: 7. 

Poland: 7. 

Portugal: 7. 

Puerto Rico: 8. 

Queensland: 8. 

Romania: 8. 

Russia: 8, 83. 

Sabah: 8. 

Samoa: 8. 

Saxony: 8. 

South Africa: 8, 136. 

Spain: 7, 8. 

Sri Lanka: 8. 

Sweden: 8. 

Switzerland: 8. 

Taiwan: 8. 

Tanzania: 8. 

Thailand: 8, 136. 

Trinidad and Tobago: 8. 

Turkey: 8. 

Uganda: 8. 



I 



( 
United Kingdom: 6, 57, 58, 110, 135, 136. ( 

( 
I 
l 
I 



192 

United States: 8, 83. 

Uruguay: 8. 

U.S.S.R.: 8, 134. 

Venezuela: 8. 

Vietnam: 8. 

Yugoslavia: 8. 
Germplasm Screening: 121, 122, 128-131. 
Glucosinolate Effect: 109, 110. 
Growth Substances: 119, 120. 

Haustoria: 20, 23-25, 29-39, 45, 64, 65, 73-76, 80, 90, 91, 99, 102, 120. 
Homoihallism: 60, 63, 130. 
Heterothallism: 60, 63, 130. 
Histopathology: 90, 91. 
Horse Radish: 6-9, 88. 
Host Differentials: 56, 57. 

Host-Pathogen Interaction: 25, 31, 69, 73, 90, 92, 122. 
Host-Pathogen Recognition System: 106, 107. 
Host Penetration: 25, 73, 74, 76, 90, 99. 
Host Range: 6, 7, 9, 10, 14, 56-59. 
Host Resistance: 99, 142. 
Host Response and Reaction: 45, 58, 59. 
Hydrogen Ion Concentration (pH): 69, 120. 



193 
Hypertrophy (Malformation): 4, 5, 11, 17, 25, 63, 64, 69, 73, 74, 83, 87, 88, 92, 137, 147. 

Hyphae: 25, 29. 

Immune: 55, 99, 105. 

Inoculum Concentration: 73. 

Inoculum Dose Relationship: 73. 

Insecticide Spray: 88. r 

Integrated Disease Management: 142. 

Isolate: 51-64, 99-102, 107-111, 119, 120, 130. 

Knop's Medium, Growth on: 64, 120. 

Kohlrabi: 6-9. 

Leaf Wetness: 76, 83, 85, 86, 90, 91. 

Lepidium: 8, 67, 142. 

Light: 69, 83. 

Lignin / Lignification: 107, 111-116. 

Local Infection: 4. 

Maledmia Africana: 7. 

Matthiola Species (Stocks): 6, 7, 17, 24. 

Metabolic Change: 92-98, 103, 104. 

Mildew: 4. 

Mixed Infection: 5, 83, 88, 89. 

Mustard: 5-7, 9, 83-91, 137, 143, 146-150. 

Mustard Mosaic Virus: 89. 



194 
Mycelium: 20, 23. 
Nutrition: 82, 83, 88, 134. 
Oilseed Rape: 6, 7, 9, 57, 61, 99-104, 122. 
Oospores: 24, 25, 60, 63, 64, 74, 130. 
Oospore Germination: 69. 
Paraperonospora: 4. 
Pathogen: 18. 

Pathogenesis: 57, 69, 73, 74, 90, 93, 99. 
Pathotype: 57, 58, 105. 
Peronospora brassicae: 20, 51. 

Peronospora parasitica: 4-6, 9, 11, 13, 14, 18, 24, 26-28, 30, 32-151. 
Peronospora parasitica -Albugo Candida, Mixed Infection: 5, 11. 
Peronospora ramicis: 20. 
Peronospora species: 5, 6, 9. 
Peroxidase: 74, 104. 
Perpetuation: 63-65. 
Phenolic Compounds: 109, 110. 
Phylogeny: 20, 21. 

Physiologic Specialization: 51-60, 145. 
Planting Time: 93, 98, 134. 

Radish: 7, 8, 23, 51, 55, 56, 63, 69, 73, 88, 99, 105, 107, 120, 135, 137, 138, 141, 143. 
Rainfall (Water): 74, 76, 83. 



195 3 

Rape: 6, 7, 9, 55, 56, 119. 

Rapeseed-Mustard: 5, 7, 9, 64, 83, 92, 134, 135, 141. 

1 

A 
Raphanus raphanistrum: 51, 55. 

Raphanus sativus: 6, 7, 8, 9, 51, 55-59, 67, 114, 118, 132. 

I 

Raphanus sativus L. var. hortensis f. minowase (Japanese Radish): 20, 45, 47-50, 99, 111- 

114, 120, 124, 125. 

Relative Humidity: 65-70, 74, 76, 81, 83, 92, 133. 

% 
Reproduction and Reproductive Structures: 20, 83. 

I 

Resistant: 17, 18, 55, 57, 99, 105, 106. 

Resistance: 17, 51, 99, 106-110, 114. 

1 

Rutabaga: 51. Q 

4 
Seed Borne: 63-65, 132. m 

I 

Seed Transmission: 63-65, 67, 74. m 

Seed Treatment: 134, 135, 138, 148-150. 

3 
Sexual Phase: 24. ,. 

Sinapis species (White Mustard): 6, 8, 51, 55. g 

9 
Sinapis alba: 6, 51, 55, 56. g 

9 
Sisymbrium species: 56. g 

9 
Soil Borne: 74. _ 

9 
Soil Treatment: 135. - 

9 
Sources of Resistance: 114, 117, 118. m 

I 

Sporangia: 91. 4 

* 
I 



196 
Sporangiophore: 91. 
Sporulation: 69, 72, 76, 89, 99, 101, 133. 
Stock: 6-9, 17, 88, 142. 
Stomata: 74, 76, 90, 99, 105. 
Susceptible: 17, 18, 51, 55, 59, 99, 105, 106. 
Swede: 6, 7, 9, 119, 120. 
Symptoms: 4, 9-17, 81, 99. 
Systemic Acquired Resistance: 106. 
Systemic Infection: 4, 11, 127. 
Taxonomy and Morphology: 20. 
Techniques: 116-134. 

Temperature: 24, 64-67, 69-74, 76, 79-81, 83-87, 89, 92, 133. 
Temperature Effect (Conidia/Conidiophores): 24, 64-67, 69-73, 133. 
Temperature Effect (Disease): 74, 84-87, 89, 92. 
Temperature Effect (Oospores): 60, 63, 64, 69. 
Toria: 6, 7, 9. 
Turnip: 6-9, 23, 51, 56, 73. 
Turnip rape: 6, 7, 9. 
Ultra Structures: 25, 26. 
Wallflower: 6, 8, 9, 17, 88. 
Watercress: 6, 9. 
White Rust: 5, 83, 87, 88, 92, 93, 98, 147. 



197 
Yield Loss: 5, 9, 137. 

Yield Increase: 137, 144, 146. 



CANADIAN AGRICULTURE. LIBRARY 



BIBLIOTHEQUE CANADIENNE DE L AGRICULTURE 

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